Polyelectrolyte multilayer films at liquid-liquid interfaces and methods for providing and using same

ABSTRACT

The present invention is directed to methods for providing a polyelectrolyte multilayer film at a liquid-liquid interface. Such methods include steps of sequentially-depositing layers of cationic and anionic polyelectrolytes at a liquid-liquid interface that is formed between immiscible first and second liquids whereby a polyelectrolyte multilayer film is provided at the liquid-liquid interface. In certain preferred embodiments, the first liquid is an aqueous solution and the second liquid is a liquid crystal. In alternative embodiments, the first liquid is an aqueous solution and the second liquid is an oil. The invention further encompasses polyelectrolyte multilayer films provided by the disclosed methods as well as applications utilizing such materials.

CROSS-REFERENCE TO RELATED APPLICATION

The present utility patent application claims the benefit of U.S.Provisional 60/697,432, filed Jul. 8, 2005, which is incorporated byreference herein in its entirety.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH

This invention was made with United States government support awarded bythe National Science Foundation—Grant Nos. CTS-0327489 and DMR-0079983.The United States has certain rights in this invention.

FIELD OF THE INVENTION

This invention relates generally to methods of functionalizingliquid-liquid interfaces. More particularly, the present invention isdirected to polyelectrolyte multilayer films formed at liquid-liquidinterfaces and methods for providing and using the same.

BACKGROUND OF THE INVENTION

The layer-by-layer technique of sequential adsorption of oppositelycharged polymers and nanoparticles onto solid surfaces has beendemonstrated in the past to be a simple and versatile method for thefabrication of supported thin films. The method creates polyelectrolytemultilayer (PEM) films by alternately immersing the surface of a solidinto solutions of polycations or polyanions. The layer-by-layertechnique can be used to deposit many types of polymers, molecules andparticulates onto surfaces, including synthetic, linearpolyelectrolytes; dendrimers; charged biomolecules such aspolynucleotides, proteins and polysaccharides; or polyvalent smallmolecular weight organic compounds. The diverse nature of thesematerials (including nanoparticles) has made possible the use of themethod in the fabrication of ion-selective membranes, chemical sensors,systems for drug and gene delivery, and patterned surfaces. It has alsobeen demonstrated that it is possible to incorporate non-ionic polymersinto multilayer films by the layer-by-layer method, and such non-ionicpolymers fall within the scope of materials that can be deposited atliquid-liquid interfaces by the methods described in this invention.

The process of PEM formation and the physical properties of theresulting films (e.g., morphology, thickness, layer interpenetration)depend on the deposition procedure, the chemical structure and molecularweight of the polyelectrolytes, and the ionic strength and pH of thedeposition solution. PEM films are most often prepared on flat solidsubstrates, but have also been formed on suspended colloidal particlesand the surfaces of macroscopic three-dimensional objects. Methods offorming PEM films on solids have typically used either (i) solids withhydrophobic interfaces in conjunction with a polyelectrolyte thatpartitions on hydrophobic substrates or (ii) solids with chargedsurfaces to initiate PEM film formation.

In contrast, the preparation of PEM films at interfaces betweenliquid-liquid phases is largely an unexplored field. However, there arevarious reasons why preparation of PEM films at liquid-liquidinterfaces, if possible, would be of great industrial value. Forexample, in the context of aqueous-liquid crystal interfaces, theformation or reorganization of PEM films overlying a liquid crystal mayresult in ordering transitions in the liquid crystal thereby providing afacile means to amplify changes in the structure of PEM films intooptical or electrical signals. Second, formation or reorganization ofPEM films at a liquid-liquid interface may provide a general andversatile approach for adding functionality to liquid crystals for useas chemical and biological sensors or as materials on which biologicalcells can be cultured. These propositions build from the observationthat the orientations assumed by liquid crystals near interfaces (the“anchoring” of the liquid crystal) are known to be highly sensitive tothe nature of the interactions between the mesogens forming the liquidcrystal and a confining interface. Depending on the structure of theinterface, the liquid crystal may align normal to the interface(homeotropic anchoring), parallel to the interface (planar anchoring),or at an angle relative to the interface (tilted anchoring). Otherorientational orderings of liquid crystals near interfaces are alsoknown.

Past studies have reported on the influence of surfactants on theorientations of liquid crystals when the surfactants are adsorbed atinterfaces of aqueous phases and thermotropic liquid crystals inemulsions (Drzaic, Liquid Crystal Dispersions. Series on LiquidCrystals; World Scientific: Singapore, 1995; Poulin et al. Science 1997,275, 1770; Mondain-Monval et al. Eur. Phys. J. B 1999, 12, 167). Morerecently, planar interfaces between thermotropic liquid crystals andaqueous solutions have been used to investigate the orientations ofliquid crystals decorated with surfactants (Brake et al. Langmuir 2002,16, 6101; Brake et al. Langmuir 2003, 16, 6436; Brake et al. Langmuir2003, 21, 8629), lipids (Brake et al. Science 2003, 302, 2094; Brake etal. Langmuir 2005, 21, 2218), and proteins (Brake et al. Science 2003,302, 2094.).

The formation of PEMs at liquid-liquid interfaces may also be used tomechanically stabilize the interface or to immobilize agents such ascatalysts of reactions at the interface. For example, if an enzyme isincorporated into a PEM at a liquid-liquid interface then the substratesand products of the enzymatic reaction could be delivered to and fromthe enzyme via either side of the PEM at the liquid-liquid interface. Inaddition, systems containing multiple enzymes could be hosted withinPEMs formed at liquid-liquid interfaces. The capacity of the PEM to hostthe enzyme could be substantially greater than is possible when enzymesadsorb directly at liquid-liquid interfaces. In addition, themicroenvironment of the enzyme can be controlled by the structure of thePEM, thus maximizing the activity and stability of the enzyme. Theformation of PEMs at liquid-liquid interfaces could also be a generaland facile route to the fabrication of free standing PEM structures whenthe liquids are chosen to be easily removed from the PEM. PEMs formed atliquid-liquid interfaces could also be used to prevent the adsorption ofbiomolecules and other molecules at liquid-liquid interfaces, thuspreventing the fouling of the interface. PEMs formed at liquid-liquidinterfaces may also change the rheological properties of the interfaces,which could find use in stabilizing emulsions and other dispersed liquidphases used in cosmetic formulations, drug delivery and othertechnologies of value to society.

It can therefore be appreciated from the foregoing that fabrication ofPEM films in combination with liquid-liquid interfaces, if possible,would yield valuable materials with utility in a wide variety ofapplications.

SUMMARY OF THE INVENTION

In a first embodiment, the present invention is directed to methods forproviding a polyelectrolyte multilayer film at a liquid-liquidinterface. Such methods include steps of sequentially-depositing layersof cationic and anionic polyelectrolytes at a liquid-liquid interfacethat is formed between immiscible first and second liquids whereby apolyelectrolyte multilayer film is provided at the liquid-liquidinterface. In certain preferred embodiments, the first liquid is anaqueous solution and the second liquid is a liquid crystal. Inalternative embodiments, the first liquid is an aqueous solution and thesecond liquid is an oil. As well, one of the liquids may be in the formof a droplet such that the polyelectrolyte multilayer film is formed atthe interface of droplet and surrounding liquid.

In another embodiment, the invention is directed to a polyelectrolytemultilayer film positioned at a liquid-liquid interface between twoimmiscible liquids. Such PEM films include sequentially-deposited layersof cationic and anionic polyelectrolytes wherein the polyelectrolytemultilayer film is positioned between immiscible first and secondliquids. In preferred embodiments, the first liquid is an aqueoussolution and the second liquid is a liquid crystal. In alternativeembodiments, the first liquid is an aqueous solution and the secondliquid is an oil. In certain other embodiments, the polyelectrolytemultilayer film contains non-ionic polymers in addition to the cationicand anionic polyelectrolytes.

In yet another embodiment, the invention encompasses a method forproviding a polyelectrolyte multilayer film at an aqueous-liquid crystalinterface. Such a method includes steps of sequentially-depositinglayers of cationic and anionic polyelectrolytes at an aqueous-liquidcrystal interface that is formed between an aqueous phase and a liquidcrystal phase whereby a polyelectrolyte multilayer film is provided atthe aqueous-liquid crystal interface. In preferred methods, thepolyelectrolyte multilayer film includes an excipient capable ofinteracting with an analyte present in the aqueous solution therebycausing a change in orientational ordering of the liquid crystal. Inparticularly preferred embodiments, the excipient is a ligand or areceptor capable of selectively-binding the analyte. Alternatively, theexcipient is a molecule capable of undergoing a chemical reaction in thepresence of the analyte.

Certain embodiments are characterized by the polyelectrolyte multilayerfilm being deposited directly on the liquid crystal phase.Alternatively, methods according to the invention may include theadditional step of seeding the aqueous-liquid crystal interface with alipid comprising a charged head group wherein deposition of thepolyelectrolyte multilayer film is facilitated by the lipid. As well,the liquid crystal may be in the form of a droplet such that thepolyelectrolyte multilayer film is formed at the interface of liquidcrystal droplet and surrounding aqueous phase.

Other embodiments of the invention encompass a modified aqueous-liquidcrystal interface. Such an interface includes: (a) an aqueous phase; (b)a liquid crystal phase; and (c) a polyelectrolyte multilayer filmpositioned between the aqueous phase and the liquid crystal phase. Suchembodiments may, optionally, encompass additional non-ionic polymers inthe multilayer film formed at an aqueous-liquid crystal interface. Suchmaterials may be formed by sequential exposure of the interface tosolutions containing non-ionic polymers.

Yet other embodiments of the invention are directed to a modified liquidcrystal comprising a liquid crystal layer and a polyelectrolytemultilayer film deposited on the liquid crystal layer. Thepolyelectrolyte multilayer film is deposited directly on the liquidcrystal layer or, alternatively, a lipid having a charged head group ispresent at the interface that facilitates formation of the PEM film. Inpreferred embodiments, the polyelectrolyte multilayer film includes anexcipient capable of interacting with an analyte present in an aqueousphase contacted with the polyelectrolyte multilayer film, theinteracting causing a change in the orientational ordering of the liquidcrystal layer. In particularly preferred embodiments, the excipient is aligand or a receptor capable of selectively-binding the analyte.Alternatively, the excipient is a molecule capable of undergoing achemical reaction in the presence of the analyte. In another embodiment,the excipient is an enzyme substrate capable of being transformed by anenzymatic analyte. Alternatively, in yet another embodiment, theexcipient is an enzyme that can catalyze the transformation of ananalyte. As well, the liquid crystal may be in the form of a dropletsuch that the polyelectrolyte multilayer film is deposited on thedroplet.

In yet another embodiment, the invention provides a method of detectingan analyte contained in an aqueous solution. Such a method includessteps of: (a) contacting an aqueous solution containing an analyte witha polyelectrolyte multilayer film deposited on a liquid crystal; and (b)determining whether a change in orientational ordering of the liquidcrystal occurs as the aqueous solution containing the analyte iscontacted with the polyelectrolyte multilayer film deposited on theliquid crystal. The presence of the analyte in the aqueous solution isindicated by the change in the orientational ordering of the liquidcrystal.

In preferred detection methods, the polyelectrolyte multilayer filmincludes an excipient capable of interacting with an analyte present inthe aqueous phase contacted with the polyelectrolyte multilayer film,the interacting causing a change in the orientation of the liquidcrystal layer. In particularly preferred embodiments, the excipient is aligand or a receptor or an enzyme capable of selectively-binding ortransforming the analyte. Alternatively, the excipient is a moleculecapable of undergoing a chemical reaction in the presence of theanalyte. In another embodiment, the excipient is an enzyme substratecapable of being transformed by an enzymatic analyte. Alternatively, inyet another embodiment, the excipient is an enzyme that can catalyze thetransformation of an analyte.

In yet a further embodiment, the invention encompasses a liquid crystaldevice, comprising: (a) a container; (b) a liquid crystal disposedwithin said container; and (c) a polyelectrolyte multilayer filmdeposited on a surface of the liquid crystal.

In another embodiment, the invention encompasses a method for providingan unsupported PEM film, the method including steps of (a) providing aliquid-liquid interface between immiscible first and second liquids; and(b) sequentially-depositing layers of cationic and anionicpolyelectrolytes at the liquid-liquid interface whereby apolyelectrolyte multilayer film is provided at the liquid-liquidinterface; and (c) removing the first and second liquids to provide anunsupported polyelectrolyte multilayer film. In certain embodiments, oneof the liquids is provided in the form of a droplet and the unsupportedpolyelectrolyte multilayer film formed by the method is in the generalshape of a hollow sphere or capsule. A particularly preferred methodincludes steps of: (a) preparing an aqueous-liquid crystal interfacebetween an aqueous phase and a liquid crystal phase; (b) depositingalternating layers of cationic and anionic polyelectrolytes at theaqueous-liquid crystal interface whereby a polyelectrolyte multilayerfilm is provided at the aqueous-liquid crystal interface; and (c)removing the aqueous phase and the liquid crystal phase to provide anunsupported polyelectrolyte multilayer film.

Other objects, features and advantages of the present invention willbecome apparent after review of the specification, claims and drawings.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 A) Schematic illustration of the geometry for producing planarinterfaces between aqueous phases and immiscible thermotropic liquidcrystals. An example of the director profile in the liquid crystal filmfor planar anchoring is also shown. The thicknesses of the slide, grid,and PEM film are not to scale. B) Structures of the liquid crystal 5CBand polyelectrolytes PSS, PAA, and PAH, referred to herein.

FIG. 2. Procedure for preparation of PEMs at an aqueous-LC interface.Method 1 (left) yields PEMs directly on the liquid crystal, while Method2 (right) incorporates a monolayer of charged lipid between the liquidcrystal and the PEM thereby facilitating deposition of polyelectrolytesthat do not partition to the aqueous-liquid crystal interface.

FIG. 3. Evidence for the formation of PEMs at aqueous-liquid crystalinterfaces. A) Quantification of the linear buildup of PEM layers usingthe fluorescence intensity of FITC-labeled PAH. B-C) Optical images(crossed polars) of 5CB in contact with water before (B) and after (C)deposition of 10 PSS/PAH bilayers directly on the aqueous-5CB interface.D) Optical image (crossed polars) of 5CB placed in water andsubsequently dried in air, for comparison to E. E-F) Optical (crossedpolars, E) and fluorescent (F) images of 10 PSS/PAH bilayers formed atthe aquous-5CB interface and subsequently dried in air. G-H) Brightfield(G) and fluorescent (H) images of 10 PSS/PAH bilayers formed at theaquous-5CB interface that has been dried in air, placed in ethanol toremove the 5CB, and dried, demonstrating freestanding PEM films. I-J)Optical images (crossed polars) of 5CB coated with a monolayer of DLEPC(I) and after deposition of 10 PAA/PAH bilayers on the DLEPC-decoratedinterface (J). Scale bars for all images are 300 μm.

FIG. 4. PEM films mediate interactions between surfactants and liquidcrystals. A-F) Optical images (crossed polars) of an uncoatedaqueous-5CB interface (A-C) and an aqueous-5CB interface coated with tenPSS/PAH bilayers (D-F) in water (A, D), 30 s after exposure to 5 mM SDS(B, E) and after subsequently exchanging the solution for an SDS-freesolution (C, F). Image in C) is after 30 s, image in F) is after 12 h.G-L) Optical images (crossed polars) of an uncoated aqueous-5CBinterface (G-I) and an aqueous-5CB interface coated with ten PSS/PAHbilayers (J-L) in water (G, J), 30 s after exposure to 5 mM DTAB (H, K),12 h after exposure to 5 mM DTAB (L), and 30 s after subsequentlyexchanging the solution for a DTAB-free solution (I). Scale bars are 300μm.

FIG. 5. Fluorescence images of A) silicon oil after deposition of 3bilayers of PSS/PAH (no fluorescently-labeled polymer) are shown and, aswell, B) silicon oil after deposition of 4 bilayers of PSS/PAH, thefinal PAH layer incorporating a fluorescently-labeled PAH polymer.

FIG. 6. Fluorescence images depicting 5 bilayers of PSS/PAH (twofluorescently-labeled PAH layers) at A) aqueous-5CB and B) aqueous-TL205interfaces.

FIG. 7. a) Schematic illustration of the experimental geometry used toprepare planar interfaces between aqueous phases and immisciblethermotropic liquid crystals. PEMs form at the LC-aqueous interface andthe LC shows planar anchoring at such interfaces. b) Structures of themolecules used in this work: the liquid crystal 5CB, repeat units ofPSS, PAA, PAH, FITC-PAH and Methacryloxyethyl thiocarbamoyl rhodamine Bcopolymerized with PSS.

FIG. 8. Quantification of the growth of PSS/PAH multilayers at the5CB-aqueous interface: Average increase in fluorescence intensity of (A)Methacryloxyethyl thiocarbamoyl rhodamine B labeled PSS for PSS-Rh/PAR(pH=8) PEMs and of (B) FITC labeled PAH measured for PSS/FITC-PAH (pH=8)PEMs grown at the 5CB-aqueous interface. Linear fits show the lineargrowth of the fluorescence intensity in both the cases.

FIG. 9. Comparison of growth of PSS/PAH PEMs grown at the 5CB-aqueousinterface Vs hydrophobic and hydrophilic solid substrates: Averageincrease in fluorescence intensity of FITC labeled PAH measured forPSS/FITC-PAH (pH=8) films grown at the 5CB-aqueous interface, cleanglass and OTS coated glass. To get an estimate for the films thicknesson 5CB, the right side Y-axis indicates the average increase inellipsometric thickness of PSS/PAH (pH=8) films grown on Si wafer.

FIG. 10. pH dependence growth of PEM of PSS/FITC-PAH on hydrophilicsilicon substrate, hydrophobic OTS coated silicon and 5CB-aqueousinterface: Average increase in fluorescence intensity of FITC labeledPAH measured for PSS/FITC-PAH (pH=5 or 8) at 5CB-aqueous interface andOTS coated glass

FIG. 11. (a) Quantification of the growth of FITC-PAH/PAA PEMs at the5CB-aqueous interface, hydrophilic native oxide glass and hydrophobicOTS coated glass: Average increase in the fluorescence intensity of FITClabeled PAH measured for FITC-PAH(7.5)/PAA(3.5) PEM and forFITC-PAH(6.5)/PAA(6.5) PEM grown at the 5CB-aqueous interface,hydrophilic native oxide glass and hydrophobic OTS coated glass for 1mg/ml concentrations of FITC-PAH.

FIG. 12. Variation of zeta potential with pH of a 5CB-aqueous interface.

FIG. 13. Average increase in ellipsometric thickness of PSS/PAR films(pH=8) grown on hyrdophilic silicon substrate and OTS coated siliconwafer.

FIG. 14. Ellipsometric thickness of PSS/FITC-PAH films (pH=5 or 8) grownon OTS coated silicon & hyrdophilic silicon substrate.

FIG. 15. Average increase ellipsometric thicknesses ofFITC-PAH(7.5)/PAA(3.5) and FITC-PAH(6.5)/PAA(6.5) PEM on hydrophilizedsilicon substrate at two different concentration of 0.2 mg/ml and 1mg/ml of FITC-PAH used for growing multilayer.

FIG. 16. (a) Quantification of the growth of FITC-PAH/PAA PEMs at the5CB-aqueous interface and hydrophilic native oxide glass: Averageincrease in the fluorescence intensity of FITC labeled PAH measured forFITC-PAH(7.5)/PAA(3.5) PEM and for FITC-PAH(6.5)/PAA(6.5) PEM grown atthe 5CB-aqueous interface and hydrophilic native oxide glass for 0.2mg/ml concentrations of FITC-PAH.

FIG. 17. Schematic illustration of a PEM supported on a lipid-decoratedsurface of a LC. Transport of the enzyme across the PEM is reported bythe change in orientation of the LC that accompanies the enzymaticdegradation of the lipid.

FIG. 18. Schematic illustration of a PEM formed using an enzyme as onecomponent: upon exposure of the PEM to a vesicle, the enzyme cleaves thecomponents of the vesicle and the flux of products of the enzymaticdegradation diffuse to the surface of the LC.

FIG. 19. Schematic illustration of reporting of botulinum toxin usingPEMs and LCs. The products of the reaction catalyzed by BoNT/A canpermeate the PEM leading to orientational changes in the LCs.

FIG. 20. Chemical structure of the repeating sequence in HA.

FIG. 21. (a) pH dependence of the ζ-potential of uncoated LC emulsions.(b) ζ-potential of (PAH/PSS)-coated 5CB-PSS and 5CB-DLEPC emulsions as afunction of layer number. The ζ-potential measurements were taken inwater (pH ˜5.7). Hollow and filled symbols indicate the PSS and PAHadsorption steps, respectively. The error in ζ-potential measurements is±5 mV.

FIG. 22. (a) Fluorescence intensity of (PAH-FITC/PSS)-coated 5CB-PSS and5CB-DLEPC emulsions as a function of layer number, as measured by flowcytometry. The measurements were taken after deposition of every bilayer(after every PAH-FITC layer). (b) Fluorescence image of the 5CB-PSSemulsion coated with seven bilayers of PAH-FITC/PSS. Scale bar is 10 μm.

FIG. 23. Director profiles and optical microcopy images of (a) 5CB-H2Oemulsions and (b) 5CBDLEPC emulsions. Scale bars are 2 μm.

FIG. 24. Cross-polarized images of (a) 5CB-PSS emulsions coated with(PAH/PSS)5 and (b) 5CBDLEPC emulsions coated with (PSS/PAH)5. Scale barsare 5 μm.

FIG. 25. (a) TEM images of hollow capsules obtained from 5CB-PSSemulsions coated with (PAH/PSS)5 and (b) an AFM image of a hollowcapsule obtained from 5CB-DLEPC emulsions coated with (PSS/PAH)5/PSS.Z-scale: 245 nm.

FIG. 26. Cross-polarized images of 5CB-PSS emulsions coated with(PAH/PSS)7 after exposure to 5 mM SDS. Scale bar is 10 μm.

DETAILED DESCRIPTION OF THE INVENTION I. In General

Before the present materials and methods are described, it is understoodthat this invention is not limited to the particular methodology,protocols, materials, and reagents described, as these may vary. It isalso to be understood that the terminology used herein is for thepurpose of describing particular embodiments only, and is not intendedto limit the scope of the present invention which will be limited onlyby the appended claims.

It must be noted that as used herein and in the appended claims, thesingular forms “a”, “an”, and “the” include plural reference unless thecontext clearly dictates otherwise. As well, the terms “a” (or “an”),“one or more” and “at least one” can be used interchangeably herein. Itis also to be noted that the terms “comprising”, “including”, and“having” can be used interchangeably.

Unless defined otherwise, all technical and scientific terms used hereinhave the same meanings as commonly understood by one of ordinary skillin the art to which this invention belongs. Although any methods andmaterials similar or equivalent to those described herein can be used inthe practice or testing of the present invention, the preferred methodsand materials are now described. All publications and patentsspecifically mentioned herein are incorporated by reference for allpurposes including describing and disclosing the chemicals, instruments,statistical analysis and methodologies which are reported in thepublications which might be used in connection with the invention. Allreferences cited in this specification are to be taken as indicative ofthe level of skill in the art. Nothing herein is to be construed as anadmission that the invention is not entitled to antedate such disclosureby virtue of prior invention.

As used herein, the term “polyelectrolyte” shall mean a polymericsubstance, either natural (e.g., protein, nucleic acid, or carbohydrate)or synthetic (e.g., poly(allylamine hydrochloride or poly(acrylicacid)), containing ionic or partially charged constituents being eithercationic or anionic.

The term “liquid crystal”, as used herein, refers to an organiccomposition in an intermediate or mesomorphic state between solid andliquid. Suitable liquid crystals for use in the present inventioninclude, but are not limited to, thermotropic, lyotropic, chromonic,smectic, nematic, ferroelectric and cholesteric liquid crystals. Theliquid crystals used in the scope of the invention may also incorporatenanoparticles such as, e.g., metallic nanoparticles.

The term “polyelectrolyte multilayer films” or “PEM films”, as usedherein, shall refer to films having at least one “bilayer” of depositedpolyanion and polycation. The term “bilayer”, as used herein, shallrefer to the accumulated layers of material deposited on a surface as aresult of having passed through at least one complete cycle of thegeneral methodologies described below and schematically shown in, e.g.,FIG. 2. Use of the term “PEM film” or “bilayer” herein is not intendedto place a restriction on the types of structures that are formed as aresult of having passed through at least one complete cycle of thegeneral methodologies described below and schematically shown in, e.g.,FIG. 2 and described in this disclosure. The term “bilayer” shall referto the sequential exposure of the interface to separate solutions of twopolyelectrolytes. It is widely understood by those skilled in the artthat the sequential exposure of an interface to polyelectrolytes ofopposite charge can lead to a range of interfacial structures and thatin some cases there is substantial mixing of the PEM with thepolyelectrolyte in solution to which the PEM is exposed. In some cases,the growth of the PEM occurs linearly with the number of cycles ofexposure, in other cases so-called exponential growth regimes areobserved. Preferred embodiments utilize films having at least twobilayers of polyanion and polycation although specific applications willdictate the optimum number of bilayers to be determined by no more thanroutine experimentation.

The following abbreviations are used throughout the present disclosure:

LbL, layer-by-layer; PEM, polyelectrolyte multilayer; PSS,poly(sodium-4-styrenesulfonate); PAH, poly(allylamine hydrochloride);PAA, poly(acrylic acid); FITC-PAH, fluorescein isothiocyanate-labeledpoly(allylamine hydrochloride; 5CB, 4′-pentyl-4-cyanobiphenyl; DLPS,1,2-Dilauroyl-sn-glycero-3-[phospho-L-serine]; DLEPC,1,2-dilauroyl-sn-glycero-3-ethylphosphocholine; SDS, sodium dodecylsulfate, DTAB, dodecyltrimethylammonium bromide.

II. The Invention

The preparation of PEMs at liquid-liquid interfaces according to theinvention involves (1) preparing or providing an interface betweenimmiscible first and second liquid phases, and (2) depositingalternating layers of polycationic and polyanionic polymers at theliquid-liquid interface. Methods of PEM formation at liquid-liquidinterfaces will now be described by reference to an aqueous-liquidcrystal example. However, it should be realized that the methodsaccording to the invention are not to be limited to any one particularliquid-liquid combination (e.g., aqueous-liquid crystal) but areapplicable to PEM formation at wide variety of liquid-liquid interfacesformed between immiscible first and second liquids. Referring now toFIG. 1A, a suitable interface between an aqueous phase and a liquidcrystal is depicted where liquid crystal is hosted in the pores of agold transmission electron microscope (TEM) grid sitting atop a glasssubstrate. The present example employs an approximately planar interfacebetween a representative liquid crystal, in this case, the nematicliquid crystal 4′-pentyl-4-cyanobiphenyl (5CB), and an aqueous solution.This assembly is prepared as follows: Glass slides are treated withoctadecyltrichlorosilane (OTS). Gold specimen grids are placed onto thesurface of glass, and liquid crystal was dispensed onto each grid andthe excess liquid crystal removed with a syringe to produce anapproximately planar surface. This geometry permits easy observation andinterpretation of the orientations of the liquid crystals at theaqueous-5CB interface and enables the sequential contact of aqueoussolutions containing polyelectrolytes of opposite charge (e.g., PSS, PAAand PAH; respective structures for these compounds shown in FIG. 1B),leading to the formation of PEM films.

The formation of PEMs at aqueous-liquid crystal interfaces then proceedsby one of two alternative methods. As one of skill in the art willappreciate after review of this disclosure, the two alternativeapproaches are certainly applicable to PEM formation at otherliquid-liquid interfaces including, for example, aqueous-oil interfaces.In a first exemplary method depicted at the left hand side of FIG. 2,PEMs are formed directly at the aqueous-liquid crystal interface byexchanging the water phase with a solution of polyanion, in this case,poly(sodium-4-styrenesulfonate) (PSS). The PSS solution is exchangedwith water after a suitable period of time (e.g., 15 minutes), and thewater is exchanged several times to rinse away any PSS not stronglyadsorbed to the water-liquid crystal interface. Then a solution ofpolycation, in this case, poly(allylamine hydrochloride (PAH), isintroduced and incubated for a period of time (e.g., 15 minutes). Afterrinsing with water, the process is repeated to produce PEMs possessing apre-selected number of polyanion/polycation bilayers (e.g., ten PSS/PAHbilayers). In support of broad applicability beyond the aqueous-liquidcrystal context, additional examples of PEM formation at liquid-liquidinterfaces using the methods described and claimed herein are providedin the examples section below (see, e.g., Example 7 which teaches PEMformation at an aqueous-isotropic oil interface).

The polyelectrolytes that can be used in the present invention include,but are not limited to, synthetic, linear polyelectrolytes; dendrimers;charged biomolecules such as polynucleotides, proteins andpolysaccharides; or polyvalent small molecular weight organic compounds.Exemplary polycations and polyanions useful in the formation of PEMfilms according to the invention include, but are not limited to thefollowing polymers to which ionic groups are covalently attached:polystyrenes, polyamines, polyesters, non-biodegradable polyurethanes,polyureas, poly(ethylene vinyl acetate), polypropylene,polymethacrylate, polyethylene, polycarbonates, and poly(ethyleneoxide)s. Particularly preferred polymers for use in the inventioninclude: poly(styrene sulfonate) (SPS), poly(acrylic acid) (PAA), linearpoly(ethylene imine) (LPEI), poly(diallyldimethyl ammonium chloride)(PDAC), poly(allylamine hydrochloride) (PAH); andpoly(sodium-4-styrenesulfonate) (PSS). Degradable polymers such aspolylactic acid and polyglycolic acid may also be used in the presentinvention. The polyelectrolytes can also include naturally occurringcomponents of the extracellular matrix of cells (e.g., laminin andcollagens) or synthetic polymers that incorporate peptides found inthese naturally occurring polypeptides. The polyelectrolytes can also bepeptide or synthetic substrates for enzymes such as proteinases andproteases. The polyelectrolytes that can be used in the presentinvention include organic and inorganic nanoparticles that have beenwidely demonstrated to be incorporated into PEMs formed at the surfacesof solids. For example, citrate stabilized gold nanoparticles arepolyelectrolytic materials that can be incorporated into PEMs. Otherexamples include carbon nanotubes and other inorganic nanostructureschemically treated by using methods well known to those skilled in theart (e.g., treatment with oleum) to charge the surface of thenanostructures. It has also been demonstrated that multilayer films ofnon-ionic polymers can be formed by the layer-by-layer methods usingprocedured identical to those used to form polyelectrolyte films. Thusthe methods described herein include formation of multilayer films fromnon-ionic polymers.

In an alternative method illustrated at the right hand side of FIG. 2,formation of PEMs is carried out on lipid-laden aqueous-liquid crystalinterfaces. The aqueous-liquid crystal interface can first be seededwith a lipid with a charged headgroup. The water phase is exchanged witha dispersion or solution of charged lipid, and the lipids are allowed toadsorb to the aqueous-liquid crystal interface for a period of time(e.g., 30 minutes). The lipid dispersion or lipid solution is exchangedwith water and the system rinsed before introducing a solution ofpolyelectrolyte that has the opposite charge of the lipid layer. PEMformation then proceeds as described for the first method described inthe preceding paragraph.

Various liquid crystals may be employed in liquid crystal-relatedapplications of the present invention. Examples of suitable liquidcrystals, include, but are not limited to, 4-cyano-4′-pentylbiphenyl (5CB), 7 CB, and 8 CB. A large listing of suitable liquid crystals ispresented in “Handbook of Liquid Crystal Research” by Peter J. Collingsand Jay S. Patel, Oxford University Press, 1997, ISBN 0-19-508442-X.Polymeric liquid crystals are also suitable for use in the device andmethods of the present invention. Because the devices and methods of thepresent invention include contacting the liquid crystal with aqueoussolutions, preferred liquid crystals employed in the invention should beinsoluble in water or have very limited solubility in water.Additionally, preferred liquid crystals employed in the invention shouldnot react with water. In one embodiment of the present invention, theliquid crystal deposited in the holding compartment of the substrate (ina grid cavity or in the depression in a support with a surface defininga depression) is 4-cyano-4′-pentylbipheny-1 (5 CB). Although varioustypes of liquid crystal may be employed, nematic and thermotropic liquidcrystals are preferred. However, smectic liquid crystals formed from 8CB are also suitable for use in the present invention. Suitable liquidcrystals further include smectic C, smectic C*, blue phases, cholestericphases, smectic A, and polymeric liquid crystals. In other embodimentsof the invention, the liquid crystal is a lyotropic liquid crystalformed in the aqueous phase and an oil is used that is immiscible withthe lyotropic liquid crystal. The lyotropic liquid crystal may be formedfrom nanoparticles.

It can be appreciated that a liquid crystal may be placed in one or moregrid(s) or depression(s) of a suitable substrate using varioustechniques. For example, a liquid crystal may be deposited in a grid orwell using a microliter syringe. As described above and in the Examples,a microliter capillary tube may then be used to remove excess liquidcrystal from the substrate surface. In one embodiment, a liquid crystalin a holding compartment of a substrate is heated into its isotropicphase at a temperature of about 50° C. and is then plunged into water ata temperature ranging from about 20° C. to 25° C. This methodology hasbeen found effective at removing air bubbles and excess liquid crystaland for producing suitable liquid crystal devices ready for adsorptionof a selected receptor molecule. As noted above, the liquid crystal istypically deposited into the grid or depression using a microlitersyringe. The liquid crystal may also be deposited into the grid ordepressions by first dissolving the liquid crystal in a volatile organicsolvent such as hexane, pentane, heptane, methylene chloride, orchloroform, depositing an appropriate amount of the dissolved liquidcrystal on the grid or depression, and allowing the solvent to evaporateleaving the liquid crystal in the grid. The liquid crystal may also bedeposited in the grids or depressions using microfluidic channels placedover the patterned surface or grid. A liquid crystal may then beinjected into the microfluidic channels and drawn into the grids ordepressions by capillary action or pressure-driven flow. In otheralternative embodiments, the liquid crystal is deposited into the gridby using inkjet printing (drop-on-demand) technology.

In general, the invention is directed to liquid-liquid interfacesfunctionalized by the presence of a PEM film. This functionalization isparticularly useful in the context of aqueous-liquid crystal interfaceswhere it is understood that liquid crystals near interfaces are highlysensitive to the nature of the interactions between the mesogens formingthe liquid crystal and a confining interface. Accordingly, a PEM filmmay, depending on interactions with molecules present in the aqueousphase, cause changes in the orientational ordering of the liquidcrystal. Example 5 below illustrates an example of a PEM filmselectively mediating interactions between two differing analytespresent in the aqueous phase and a liquid crystal. Specifically, Example5 demonstrates how a liquid crystal functionalized according to theinvention is caused to discern between the two surfactants, sodiumdodecyl sulfate (SDS) and dodecyltrimethylammonium bromide (DTAB),respectively, in aqueous solution. In effect, a PEM film provided at anaqueous-liquid crystal interface may tailor the selectivity of theliquid crystal in regard to the liquid crystals response to particularanalytes of interest.

As is known to those skilled in the art, changes in optical propertiesof the liquid crystal can be quantified by using optical instrumentationsuch as, but not limited to, plate readers, cameras, scanners,photomultiplier tubes. Because the dielectric properties of liquidcrystals also change with orientational order, measurements ofelectrical properties of liquid crystals can also be used to reportchanges in the interactions of molecules with liquid crystals. In someembodiments of the present invention, the optical and electricalmeasurements lead to determination of the anchoring energy of the liquidcrystal at the PEM-decorated interface. In one embodiment, thetorque-balance method is used to determine the anchoring energy.

In certain liquid crystal-related embodiments, the invention is directedto PEM films that include one or more excipients so as to, incombination with the PEM film, functionalize a liquid-liquid interface.Accordingly, excipients present in a PEM film may, depending oninteractions with each other, the PEM film, and/or molecules present inthe aqueous phase, cause changes in the orientation of the liquidcrystal.

For example, various receptor species may be combined with the PEM filmand, if an aqueous solution includes a sufficient amount of a compoundthat interacts with the receptor species, a change in the orientation ofthe liquid crystal will occur indicating the interaction (binding and/orchemical reaction) between the receptor species and the compound.Typically, the liquid crystal is viewed through polarized light todetermine whether the orientation has been altered. In one embodiment, apolarized light microscope is used and may further be used inconjunction with cross polarizers.

Examples of suitable receptor species to be combined with a PEM filminclude, but are not limited to peptides and proteins, includingintegral membrane proteins such as glycoproteins, cell signalingproteins such as G proteins, growth factor receptors, ion channelproteins, antibodies, proteoglycans, and integrins. As well, moleculessuch as hormones (e.g. estrogen, testosterone, glucagons, andepinephrine), hormone receptor proteins, growth factors, insulin,biotin, sugars (e.g. glucose, lactose), DNA, RNA, collagen,pharmaceuticals, enzyme inhibitors, peptides, polypeptides, nucleotides,oligonucleotides, antibodies, immunoglobulins, chelating agents, andmetal ions (collectively, “ligands”) may also be contained within thePEM film. Those skilled in the art will recognize that various othermolecular species may be used in accordance with the present invention.For example, enzymes or substrates for enzymes can be incorporated intoPEM films.

As directed herein, aqueous solutions containing compounds may becontacted with a PEM film including an excipient on a liquid crystal todetect interactions or chemical reactions between the compound and theexcipient. In this manner, the present invention allows one to detectinteractions (binding and/or chemical reaction) of known analytes with agiven excipient or may be used to detect or identify a given analyte inan aqueous solution. Various analytes may be determined in accordancewith the present invention. A liquid crystal device prepared using anappropriate PEM in combination with a receptor or ligand species may beused to detect eukaryotic and prokaryotic organisms, bacteria, viruses,DNA, RNA, proteins, enzymes, ions, and cells in an aqueous solution thatis circulated or passed over or contacted with the functionalized PEMfilm atop the liquid crystal. The role of the excipient and analyte maybe reversed by changing which species is hosted within the PEM layer atthe liquid crystal-aqueous interface. Examples of such interactions arebiotin and avidin, streptavidin, and antibiotin-IgG; growth factors andgrowth factor receptors; hormones and hormone receptors; enzymes andenzyme inhibitors, substrates, and initiators; antibodies and antigens;integrins and components of the extracellular matrix; cell signalingproteins as part of a cascade; and ion channel proteins and ions andactivating ligands. Generally, analyte concentrations in the aqueoussolutions may range from 1 fM to 1 M with the desirable concentrationdepending on the nature of the interaction between the analyte and thereceptor. For biological analytes, the pH of aqueous solutions shouldtypically range from 6 to 9.

In one embodiment, the invention provides a method for determining achange in the oxidation state of a molecule contained within a PEM filmoverlaying a liquid crystal. In such embodiments, the molecule includesa group such as, but not limited to, a ferrocene group that may beoxidized or reduced upon contact with an oxidizing agent, a reducingagent, an applied oxidizing potential, or an applied reducing potential.In the method, a liquid crystal device immersed in an aqueous solutionis contacted with an oxidizing agent, a reducing agent, an appliedoxidizing potential, or an applied reducing potential. In the liquidcrystal device of such methods, the molecule is contained within a PEMfilm deposited on the top surface of a liquid crystal that is located inthe holding compartment of a substrate as described above. A change inthe orientation of the liquid crystal upon contacting the liquid crystaldevice with the oxidizing agent, the reducing agent, the appliedoxidizing potential, or the applied reducing potential indicates thatthe oxidation state of the molecule has changed. Examples of groups thatmay be oxidized or reduced on the receptor molecule include, but are notlimited to, ferrocene, quinone, metal tri-nitriloactetic acid complexes,ferricyanide, viologens, metal porphyrins, alcohols, aldehydes,organosulfur compounds, anthracene, azobenzene, benzophenone,nitrobenzene, Ru(bpy)₃ ^(n+), tetracyanoquinodimethane (TCNQ),tetrathiafulvalene, and other biological redox-active species such asbut not limited to neurotransmitters. One group of suitable moleculeswith groups that may be oxidized or reduced includes(ferrocenylalkyl)trialkyl-ammonium halides such as(ferrocenylalkyl)trimethylammonium chlorides and bromides such as11-(ferrocenylundecyl)trialkylammonium bromide. The aqueous solution inwhich the liquid crystal device is immersed may also include surfactantssuch as cationic surfactants, anionic surfactants, and/or zwitterionicsurfactants. Examples include ferrocenyl surfactants;alkyltrimethylammonium halides; alkyl sulfates; phospholipids such asdilaurylphosphatidyl choline, dipalmitoylphosphatidyl choline,dilaurylphosphatidyl ethanolamine, dipalmitoylphosphatidyl ethanolamine,and combinations of these; and polymeric surfactants, such ashydrophobically modified ethylhydroxyethyl cellulose (HM-EHEC).Quaternary ammonium compounds suitable for use as surfactants include,but are not limited to, CTAB and DTAB. Typically, the aqueous solutionalso includes a salt such as, but not limited to, Li₂SO₄. Otherbuffering agents and salts may be included in the aqueous solutions suchas, but not limited to, sodium halides, potassium halides, sodiumsulfate, potassium sulfate, sodium phosphate, potassium phosphate, tris,HEPES, and MOPS.

The present methodology may be adapted to a variety of liquid crystaldevice designs. For example, the present methodology may be employed tomodify a variety of liquid crystal devices, including those described inU.S. Published Patent Application 2003/0194753 A1 to Abbott et al.,which is incorporated herein by reference. In general, liquid crystaldevices useful in the present invention will include a container havingan inlet and an outlet with the liquid crystal housed in the containeron a suitable substrate. For example, the device may include a glassslide on which a grid is positioned that is disposed inside a container.The container includes an inlet and an outlet through which a sample maybe introduced and removed from container. Inlet and outlet may beconfigured to project out the sides of container or alternatively maysimply be holes defined by side walls. Those skilled in the art willrecognize that various other configurations are possible and may beused.

Quantitative determination of kinetic parameters from the appearance ofthe liquid crystal may be accomplished by plotting a measure of theoptical texture (such as the average brightness (grayscale or an RGBchannel), standard deviation of any measure of brightness, anchoringenergy, or a Fourier transform of the image) versus time. Electricalmeasurements, as described above, can also be performed and quantified.The data may then be analyzed by fitting the data with a model of thekinetic behavior for the given interaction (e.g. a surface reactionanalogue to the Michaelis-Menton equation for enzyme kinetics).

Because the devices of the present invention may be used to detect thepresence of compounds in flowing streams, the devices may be used tocontinuously monitor the presence of a compound that interacts with thePEM film or PEM film/excipient combination on the surface of a liquidcrystal. Additionally, the devices of the present invention may be usedto monitor water quality.

In certain embodiments, the formation of PEMs at liquid-liquidinterfaces is useful to mechanically stabilize an interface or toimmobilize agents such as catalysts of reactions at the interface. Forexample, if an enzyme is incorporated into a PEM at a liquid-liquidinterface then the substrates and products of the enzymatic reactioncould be delivered to and from the enzyme via either side of the PEM atthe liquid-liquid interface. In addition, systems containing multipleenzymes could be hosted within PEMs formed at liquid-liquid interfaces.The capacity of the PEM to host the enzyme could be substantiallygreater than is possible when enzymes adsorb directly at liquid-liquidinterfaces. In addition, the microenvironment of the enzyme can becontrolled by the structure of the PEM, thus maximizing the activity andstability of the enzyme.

In other embodiments, PEMs formed at liquid-liquid interfaces are usedto prevent the adsorption of biomolecules and other molecules atliquid-liquid interfaces, thus preventing the fouling of the interface.

PEMs formed at liquid-liquid interfaces are also envisioned to changethe rheological properties of the interfaces, which finds use instabilizing emulsions and other dispersed liquid phases used in, forexample, cosmetic formulations and drug delivery. PEMs formed atinterfaces of emulsion droplets can provide means of encapsulation for,for example, protection of the encapsulant from the environment orcontrolled release of the encapsulant.

The methods of the present invention may also be used to createthree-dimensional microstructures consisting of unsupported PEM films.The formation of PEMs at liquid-liquid interfaces certainly enables ageneral and facile route to the fabrication of free standing PEMstructures when the liquids are chosen to be easily removed from thePEM. For example, the PEM film may be deposited at a liquid-liquidinterface which is then removed by dissolving one or more liquid phases,drying one or more liquid layers, or applying other suitable means toleave an unsupported PEM film. Example 4 below describes the fabricationof unsupported PEM films by drying an aqueous phase and then dissolvinga liquid crystal phase to leave the PEM film, previously deposited therebetween, in a free-standing microstructure. It will be appreciated thatmore complex microstructures could be created based on these simpleprinciples (e.g., by depositing PEMs with different electrostaticcharacters in different liquid-liquid regions and/or by iterativeadditions of subsequent structures above the deposited polymer).

The formation of unsupported PEM films prepared as described aboveprovides a general and facile method for the fabrication of nano-,micro- and milli- and larger scale structures. By incorporation ofvarious polyelectrolytes into the PEM, it is possible to make drugdelivery nano or micro devices that are transported to a tumor or woundand release active ingredients that destroy the tumor or heal the wound.Many other possible applications for such unsupported PEM films exist,including, but not limited to, sensors that are formed from asymmetricPEM structures and that change shape upon encountering a targetedanalyte. Unsupported PEM structures could also be used forchemomechanical transduction and other types of energy exchange wherethe reaction of a chemical species with one surface of the PEM leads toa mechanical deformation of the PEM against a resisting force. Otherapplications are evident to those skilled in the art of fabricatingunsupported nano- and micro-structures.

The inventors have previously shown that it is possible to prepare,e.g., layer-by-layer (LbL) films on an approximately planar interfacebetween the nematic LC 4′-pentyl-4-cyanobiphenyl (5CB) hosted in a goldgrid, and an aqueous phase (water). The LbL films preserved the planaranchoring of 5CB in water. Furthermore, the multilayer films wereobserved to mediate the interactions between solutes dissolved in theaqueous phase and the LC. One attractive feature of using LbL assemblyto modify the surface properties of LCs is the flexibility of theapproach: different materials can be assembled, including polymers,proteins, DNA, multivalent ions, and nanoparticles. The technique can bealso transferred to three-dimensional substrates, such as colloidalparticles, biomolecule crystals, macroporous membranes, and porousbeads. Recently, three-layered polymer membranes (consisting oflecithin, chitosan and pectin) were deposited on oil-in-water emulsionsof tuna or corn oil. These polymers were deposited without intermediatewashing steps and significant droplet aggregation was observed.

Accordingly, emulsions formed from LCs are useful for creation ofLC-based sensors because they have a much higher surface area than LCsformed at planar interfaces, they are mobile, and frustrated statesassumed by LCs within droplets provide an additional opportunity to tunethe response of LCs to interfacial events. Previous studies on LCemulsion droplets have shown their usefulness for studying a variety ofphenomena, including the rotational motion of particles, the effect ofhydrodynamic flows, and the effect of electric fields. Such emulsionshave been prepared by methods including photo-polymerization, dispersionpolymerization, shearing droplets and subsequent crystallizationfractionation, ultrasonication and droplet break-off in a co-flowingstream.

In an example below, the inventors illustrate how stable nematic LCemulsions are formed by sonication. In this particular exemplaryembodiment, layer-by-layer growth of multilayers of PAH and PSS onmicrometer-sized 5CB-PSS and 5CB-DLEPC emulsion droplets was confirmedwith ζ-potential measurements, flow cytometry, and fluorescencemicroscopy. The LC in the 5CB-PSS emulsion droplets assumed a bipolarconfiguration while the LC in the 5CB-DLEPC emulsion droplets adopted aradial configuration, and these configurations were preserved aftermultilayer assembly. In addition, the multilayers were used to influencethe interaction of analyte with the LC core. When the 5CB cores of themultilayer-coated emulsion droplets were dissolved with ethanol, hollowcapsules were formed. The exemplary results demonstrate that LbLassembly can be applied to the mobile interfaces between LC droplets andaqueous phases, which represent an important development in thepreparation of chemically-tailored interfaces for use inmicrometer-sized biological and chemical sensors. In addition, thisapproach is applicable to other oils, allowing a versatile route toproduce hollow polymeric capsules. Such a development offersconsiderable advantages over solid templates.

The following examples are offered for illustrative purposes only, andare not intended to limit the scope of the present invention in any way.Indeed, various modifications of the invention in addition to thoseshown and described herein will become apparent to those skilled in theart from the foregoing description and the following examples and fallwithin the scope of the appended claims.

III. EXAMPLES

The following materials and methodologies were utilized in the examplesdiscussed in greater detail below.

Materials. Poly(sodium-4-styrenesulfonate) (PSS, M_(w), 70 kDa) andpoly(allylamine hydrochloride) (PAH, M_(w), 70 kDa) were purchased fromSigma-Aldrich and used without further purification. PAA was purchasedfrom Polysciences. Fluorescein isothiocyanate-labeled PAH (FITC-PAH,M_(w), 70 kDa) was prepared as described by Caruso and coworkers. Thenematic liquid crystal 4′-pentyl-4-cyanobiphenyl (5CB) was purchasedfrom EMD Chemicals (Hawthorne, N.Y.) and used without furtherpurification. Gold specimen grids (bars 20 μm thick and 55 μm wide,spaced 283 μm apart) were obtained from Electron Microscopy Sciences(Fort Washington, Pa.) and cleaned sequentially in ethanol, methanol,and methylene chloride. The phospholipids1,2-Dilauroyl-sn-glycero-3-[phospho-L-serine] (DLPS) and1,2-dilauroyl-sn-glycero-3-ethylphosphocholine (DLEPC) were purchasedfrom Avanti Polar Lipids (Alabaster, Ala.). Sodium dodecylsulfate (SDS)was purchased from Sigma-Aldrich. SDS was purified by recrystallizationfrom ethanol. Deionization of a distilled water source was performedwith a Milli-Q system (Millipore, Bedford, Mass.) to give water with aresistivity of 18.2 MΩ·cm. Glass microscope slides were Fisher's FinestPremium Grade obtained from Fisher Scientific (Pittsburgh, Pa.).Octadecyltrichlorosilane (OTS) was obtained from Fisher Scientific.Sylgard 182 elastomer and curing agent (PDMS) was obtained from DowCorning Corp. (Midland, Mich.). 2 Ton Clear epoxy resin and hardener wasobtained from ITW Devcon (Danvers, Mass.).

Preparation of optical cells. A detailed description of the methods usedto prepare and examine the liquid crystal hosted within optical cellscan be found in Brake et al. Langmuir 2002, 16, 6101. Briefly, glassmicroscope slides were cleaned according to published procedures (Skaifeet al. Chem. Mater. 1999, 11, 612) and coated withoctadecyltrichlorosilane (OTS). The quality of the OTS layer wasassessed by checking the alignment of 5CB confined between twoOTS-treated glass slides. Any surface not causing homeotropic anchoringof 5CB was discarded. A small square of OTS-coated glass (ca. 5 mm×5 mm)was fixed to the bottom of each well of an 8-well chamber slide (NalgeNunc International, Rochester, N.Y.) with epoxy and cured overnight at60° C. The wells were rinsed several times with ethanol to removeuncured monomer and subsequently dried. Gold specimen grids that werecleaned sequentially in methylene chloride, ethanol, and methanol wereplaced onto the surface of the OTS-treated glass slides, one per well.Approximately 1 μL of 5CB was dispensed onto each grid and the excessliquid crystal was removed with a syringe.

Preparation of phospholipid solutions. The appropriate amount ofchloroform solutions of the phospholipids DLPS or DLEPC (10 mg/mL) weredispensed into glass tubes and dried under a stream of nitrogen. Thetubes were then left to dry under vacuum overnight and stored in a −20°C. freezer until needed. The lipids were reconstituted to 100 micromolarin water, hydrated overnight at room temperature, and sonicated for 30min in a bath sonicator. Upon reconstitution the lipid solutions wereclear and were used without filtering. The dried lipids were resuspendedand used within one month of drying.

Liposome preparation. Chloroform solutions of pure DLPS or DLEPC weredispensed in a glass tube and dried under a low flow of N₂ to form athin lipid film. Residual solvent was removed under vacuum at 50° C. forseveral hours. The resulting lipid film was hydrated overnight at roomtemperature (above the lipid transition temperature) with an appropriatevolume of water to yield a final lipid concentration of 100 μM. Thelipid solutions appeared clear after this step, suggesting that bothDLPS and DLEPC either dissolved or formed micellar aggregates insolution. Nevertheless, the lipid suspensions were sonicated for 30 minin a bath sonicator at room temperature to produce small unilamellarliposomes. Upon reconstitution the lipid solutions were clear and wereused without filtering.

LbL Deposition at aqueous-liquid crystal interface. PSS, PAH and PAAsolutions were of concentration 1 mg/mL in 0.1 M or 0.5 M NaCl. FITC-PAHsolutions were of concentration 0.2 mg/mL in 0.25 M or 0.5 M NaCl. PAHand PAA solutions used to form PAH/PAA multilayers were adjusted withHCl to pH 7.0; the pH was not adjusted for other solutions. Foradsorption of each polyelectrolyte at the aqueous-liquid crystalinterface, 1 mL of polyelectrolyte solution was flowed through the flowcell and incubated for 10 min (0.1 M NaCl) or 15 min (0.5 M NaCl). Three1 mL aliquots of water were then rinsed through with 1 min incubationeach to remove free polyelectrolyte from the cell. In the absence oflipid seeding, PSS/PAH multilayers were started with PSS. A PSS layerfollowed DLEPC seeding of aqueous-liquid crystal interfaces and PAHfollowed DLPS-seeding. FITC-PAH layers were followed by five 1 mL waterrinses to remove any free fluorescent molecules.

Polarized microscopy of PEMs at aqueous-liquid crystal interface. Theorientation 5CB within each optical cell was examined withplane-polarized light in transmission mode on an IX-71 invertedmicroscope with crossed polarizers. The source light intensity levelswere constant for all images of the same magnification. Homeotropicalignments of the liquid crystal were determined by observing theabsence of transmitted light regardless of rotation of the sample.Images were taken with a QImaging MicroPublisher 3.3 RTV color camera onautoexposure (unless noted) and controlled via ImagePro Expresssoftware.

Fluorescent microscopy of PEMs at aqueous-liquid crystal interface. PEMswere prepared at the aqueous-liquid crystal interface as described aboveand then imaged with an Olympus IX-71 inverted microscope using afluorescence filter cube with an excitation filter and emission filter.Images were taken with a QImaging MicroPublisher 3.3 RTV color cameracontrolled via ImagePro Express software. Unless noted, the exposuretimes for 4× and 10× images were 2 s and 500 ms, respectively. Thefluorescent images were taken with the liquid crystal/OTS glassinterface toward the objective.

EXAMPLE 1 Procedure for Preparation of PEMs at the Aqueous-LC Interface

In this example, the direct formation of a PEM film at an aqueous-liquidcrystal interface is described. PEM films were formed directly at anaqueous-5CB interface by initially contacting the surface of the 5CB for15 minutes with a solution of poly(styrene-4-sulfonate) (PSS, 1 mg/mL in0.5 M NaCl), which has been shown to partition onto hydrophobicinterfaces, and alternately exposing the interface to a solution ofpoly(allyl amine hydrochloride) (PAH, 1 mg/mL in 0.5 M NaCl). Thesesteps were repeated ten times. Initial experiments used the standardmethod of dipping the substrate (in our case, 5CB hosted in a TEM gridon a glass slide) into each solution. However, this process sometimes(but not always) displaced the 5CB from the grid upon its passagethrough the air-water meniscus. The inventors found that a more reliablemethod was to place the grid containing 5CB at the bottom of a smallwell and exchange solutions in such a way that the meniscus never fellbelow the interface of the 5CB.

To verify that a PEM film formed at the aqueous-5CB interface, one ormore layers of fluorescently-labeled PAH (FITC-PAH) were incorporatedinto the film. Fluorescence imaging showed the incorporation of FITC-PAHinto the film at the aqueous-5CB interface (data not shown). To quantifythe build-up of the PEM film, the inventors deposited three bilayers ofPSS/PAH directly at the aqueous-5CB interface followed by seven bilayersof PSS/FITC-PAH. The fluorescence intensity of the regions within thegrid squares (i.e., corresponding to regions of PEM film deposited on5CB) increased linearly with the number of FITC-PAH layers (FIG. 3A).These results confirm the formation of PEM films directly at theaqueous-5CB interface.

EXAMPLE 2 Procedure for Preparation of PEMs at the Aqueous-LC Interface

This example describes an alternative approach leading to formation ofPEM films on the aqueous-5CB interface by the seeding of the aqueous-5CBinterface with the anionic phospholipid1,2-dilauroyl-sn-glycero-3-[phospho-L-serine] (DLPS) or the cationicphospholipid 1,2-dilauroyl-sn-glycero-3-ethylphosphocholine (DLEPC)prior to introducing the initial polyelectrolyte. Phospholipids in theform of an aqueous suspension of small unilamellar vesicles havepreviously been shown to spontaneously adsorb to hydrophobic interfaces,including liquid crystals. After forming DLPS or DLEPC monolayers at theaqueous-5CB interface by incubating the interface in a solution of smallunilamellar vesicles (100 micromolar lipid in water), the inventorsexchanged the lipid solution for a lipid-free solution and subsequentlyintroduced polyelectrolyte solutions (1 mg/mL in 0.5 M NaCl) in order toform PEM films. For DLEPC-laden interfaces, which are cationic innature, the inventors first incubated the interface in a poly(acrylicacid) (PAA) solution for 15 minutes, and alternated with PAH solutionsfor the same amount of time. For anionic DLPS-laden interfaces, theinventors started with PAH and alternated with PAA. The process wasrepeated ten times and used a FITC-labeled PAH as the final PAH layer inboth cases. Fluorescent images of the 5CB indicated the presence ofFITC-PAH (data not shown), consistent with formation of PAA/PAH films atthe lipid-laden aqueous-5CB interface.

The results above are consistent with the formation of PEM films at theaqueous-5CB interface. The higher mobility of the polyelectrolytes atthe aqueous-liquid crystal interface, as compared to their mobility onthe conventional solid substrate, did not prevent the formation of thePEM film.

EXAMPLE 3 Orientational Order in the Liquid Crystal in Relation to theFormation of PEM Films

This example demonstrates that the orientational order of the liquidcrystal can be coupled to the formation of PEM films (FIG. 3B-E).Polarized light microscopy was used to characterize the orientationalorder within the liquid crystal, prepared as described in Example 1above. The optical appearance of 5CB when immersed under water (nolipids or polyelectrolyes) was bright with pale yellow-pink interferencecolors (FIG. 3B). This appearance can be explained in the followingmanner: At the 5CB-glass interface, the orientation of the liquidcrystal is anchored perpendicular to the glass due to the monolayer ofoctadecyltrichlorosilane (shown schematically in the director profile inFIG. 1). Consequently, the bright optical appearance of the liquidcrystal is the result of in-plane birefringence of the liquid crystalaligned parallel to the aqueous-5CB interface. The presence of darkbrushes emanating from points generally along the edges of eachcompartment indicates that there is a variation of the azimuthal(radial) orientation of the liquid crystal within each compartment ofthe grid. The deposition of a PEM consisting of ten bilayers of PSS/PAHled to only minor changes in the optical appearance of the liquidcrystal when viewed under water (FIG. 3C). The differences observed(changes in the position of the dark brushes, for example) were alsoobserved for controls in which water was repeatedly exchanged over the5CB. The lack of change in the optical appearance of the liquid crystalsuggests that both the process of PEM film formation and the presence ofthe PEM film itself do not significantly affect the orientations of theliquid crystal when in contact with water in this experiment. However, acoupling between the liquid crystal and PEM is evident when the PEM wasremoved from the water into air.

While the presence of a PEM film did not influence the anchoring of 5CBin contact with aqueous solutions, it was determined if the presence ofthe PEM film would be able to maintain the orientation of 5 CB if the 5CB were removed from the aqueous environment and dried in air. In theabsence of a PEM or lipid layer, 5CB showed the expected brightappearance (planar anchoring) in water (e.g., FIG. 3B), and the typicaldark appearance (homeotropic anchoring) when subsequently dried in air(FIG. 3D). In contrast, 5CB on which ten bilayers of PSS/PAH had beendeposited (formed directly at the aqueous-5CB interface) exhibited anoptical appearance that was bright, corresponding to planar anchoring ofthe 5CB (FIG. 3E). This appearance was essentially unchanged by thedrying process (compare FIGS. 3C and 3E). Fluorescence microscopyindicated a fairly uniform distribution of the top FITC-PAH layer of thePEM film on the 5CB (FIG. 3F). This result provides further evidencethat PEM films form at the aqueous-5CB interface and demonstrates therobustness of the PEM film on the 5CB. In addition, this resultindicates that a coupling exists between the orientations of 5CB andpresence of a PEM film at the 5CB surface: The PEM preserved planaranchoring of the 5CB under conditions that would normally lead tohomeotropic anchoring. Samples that were intentionally handled roughlyto dislodge regions of the PEM film from the 5CB surface showed areas ofplanar anchoring, which corresponded to regions covered by the PEM film,and regions of homeotropic anchoring, which corresponded to areas inwhich the PEM had been removed (data not shown).

The coupling between PEM film formation on lipid-seeded aqueous-5CBinterfaces and the order in the liquid crystal was also investigated.After depositing DLEPC at the aqueous-5CB interface, the opticalappearance of 5CB changed from bright (e.g., FIG. 3B) to dark (FIG. 31)within ˜1 min, indicating a change from planar to homeotropic anchoringof the 5CB. A similar transition has been observed for zwitterinicphospholipids, albeit at much longer timescales (˜2 h), and has beenshown to be the result of a coupling between adsorbed lipid and theliquid crystal. The homeotropic orientation of 5CB was maintained atleast 30 min after replacing the lipid solution with lipid-free water.The appearance of the 5CB did not change after depositing ten PAA/PAHbilayers on the DLEPC-laden interface (FIG. 3J). Similar results wereobserved when the lipid layer was DLPS rather than DLEPC (data notshown).

EXAMPLE 4 Unsupported PEM Films are Stable after Removal of SupportingLiquid Layers

This example demonstrates the PEM films are stable after removal of thesupporting liquid layers. Ten bilayers of PSS/PAH were deposited (formeddirectly at the aqueous-5CB interface) on 5CB hosted in a TEM grid, asdescribed above. The 5CB exhibited an optical appearance that wasbright, corresponding to planar anchoring of the 5CB (FIG. 3E). Next,and as described in Example 3, the 5CB was removed from the aqueoussolution and dried. Fluorescence microscopy indicated a fairly uniformdistribution of the top FITC-PAH layer of the PEM film on the 5CB (FIG.3F). After preparing PEM films on 5CB and drying the samples in air, theinventors removed the TEM grid from the OTS-glass slide and immersed thegrid-5CB-PEM sample in ethanol to dissolve the 5CB. After the ethanolevaporated, this process yielded PEM films that, while significantlydamaged, spanned many of the TEM grid squares as freestanding PEM films(FIG. 3G-H).

EXAMPLE 5 PEMs Selectively Mediate Interactions Between Analytes andLiquid Crystals

This example demonstrates a PEM film's ability to mediate interactionsbetween an analyte present in an aqueous solution and a liquid crystalupon which the PEM film is deposited. The inventors exposed aqueous-5CBinterfaces decorated with PSS/PAH multilayer films to solutions of thesurfactants sodium dodecyl sulfate (SDS) and dodecyltrimethylammoniumbromide (DTAB). In the absence of a PEM film, the adsorption of eitherSDS or DTAB to aqueous-5CB interfaces is fast (seconds) and leads to atransition from planar to homeotropic orientation of the 5CB (FIG. 4A-B,G-H). When the bulk SDS or DTAB solution is replaced with a solutionfree of surfactant both SDS and DTAB desorb from the aqueous-5CBinterface on the timescale of seconds and the 5CB orientation becomesplanar (FIG. 4C, I).

The presence of a PEM film at the aqueous-5CB interface had no apparentinfluence on the adsorption of SDS to the interface, but changed theadsorption characteristics of DTAB. When the PEM-coated interface wasexposed to 5 mM SDS, a transition from planar to homeotropic orientationof 5CB occurred on a timescale comparable to that without a PEM filmpresent (FIG. 4D-E). However, when the inventors exposed a PEM-coated5CB interface to a solution of 5 mM DTAB, they observed no transitionfrom planar to homeotropic orientation of the 5CB, even after 12 h ofexposure (FIG. 4J-L). Similar results were observed for 9.5 PSS/PAHbilayers (terminating in an anionic PSS layer, data not shown),suggesting that the phenomenon is not dependent on the electrostaticnature of the topmost PEM layer. While the PEM film showed no influenceon the kinetics of SDS adsorption to the aqueous-5CB interface, theinventors observed the desorption kinetics to depend strongly on thepresence of the PEM. The homeotropic orientation observed for 5CB coatedwith a PEM film and exposed to SDS solution was maintained for 12 hoursafter replacing the SDS solution with a SDS-free solution (FIG. 4F).

The presence of SDS creates a texture apparent in optical micrographsthat are reminiscent of wrinkled fabric (FIG. 3E, inset). This type oftexture was not observed when SDS was adsorbed to 5CB in the absence ofa PEM film (FIG. 4B, inset). The presence of DTAB did not produce anyobvious wrinkled texture of the PEM films, though there did appear to bea roughening in the appearance of the PEM-coated 5CB when exposed toDTAB for 12 h (FIG. 4L).

EXAMPLE 6 Formation of PEM Films at Liquid-Liquid Interfaces;Aqueous-Liquid Crystal Interfaces are Representative of Liquid-LiquidInterfaces

The present invention contemplates formation of PEMs at liquid-liquidinterfaces beyond aqueous-liquid crystal interfaces. One of skill in theart will appreciate that properties of aqueous-liquid crystal interfacesare representative of the broader class of liquid-liquid interfaces. Forexample, diffusion coefficients at aqueous-isotropic oil andaqueous-liquid interfaces have been demonstrated to be comparable.Fluorescence recovery after photobleaching (FRAP) measurements of thephospholipid dilauroylphosphocholine (DLPC) adsorbed at theaqueous-liquid crystal interface yielded diffusion coefficients of 6 to15×10⁻¹² m²/s for DLPC at the water-liquid crystal interface(fluorescein-dipalitoylphosphocholine probe). [Brake et al. Langmuir2005, 25:2218] Yu et al. measured DLPC diffusivity at a water-heptaneinterface to be 20×10⁻¹² m²/s (NBD probe). [Adalsteinsson, T.; Yu, H.Langmuir 2000, 16:9410.] These values demonstrate that the diffusion oflipids at aqueous-liquid crystal interfaces and aqueous-oil interfacesare comparable and substantiate the formation of PEM films across abroad range of liquid-liquid interfaces.

EXAMPLE 7 Formation of a PEM Film at an Aqueous-Isotropic Oil Interface

In order to further support the broad applicability of the presentinvention to the formation of PEM films generally at liquid-liquidinterfaces, this example describes the formation of a PEM film at theinterface between water and an immiscible silicon oil. The inventorsprepared PEM films on the aqueous-silicon oil interface usingsubstantially the same procedure previously-described in Example 1above. After repeated exposure to alternating PSS and PAH solutions, andending with a fluorescently-labeled PAH solution, a uniform brightfluorescence was observed that indicated the presence of a PEM film atthe aqueous-silicon oil interface. FIG. 5 shows fluorescence imagesdepicting this PEM formation at an oil-water interface. Specifically,FIG. 5 shows fluorescence images of A) silicon oil after deposition of 3bilayers of PSS/PAH (no fluorescently-labeled polymer) are shown and, aswell, B) silicon oil after deposition of 4 bilayers of PSS/PAH, thefinal PAH layer incorporating a fluorescently-labeled PAH polymer. Theincrease in fluorescence intensity demonstrates the formation of a PEMfilm at the aqueous-silicon oil interface. The presence of silicon oilwas verified after the treatment by observing the edges of the oil filmwith brightfield microscopy.

EXAMPLE 8 Formation of a PEM Film on the Fluorinated Liquid CrystalTL205

This example describes the formation of a PEM film on yet another liquidcrystal, namely, the fluorinated liquid crystal TL205. A procedure assubstantially described in Example 1 above was performed to provide aPEM layer at an aqueous-TL205 interface. Referring to FIG. 6, the PEMfilm was successfully formed at the aqueous-5CB interface asdemonstrated by fluorescence images depicting 5 bilayers of PSS/PAH (twofluorescently-labeled PAH layers) at A) aqueous-5CB and B) aqueous-TL205interfaces. The intensity on TL205 appears similar to that on 5CB,indicating that PEM film formation is similar on the two liquidcrystals.

EXAMPLE 9 Characterization of Growth of PEM Films at Aqueous-LCInterfaces

In this example, the inventors describe a detailed characterization ofthe growth of PEM films at aqueous-LC interfaces. The growthcharacteristics of poly(sodium-4-styrenesulfonate) (PSS)/poly(allylaminehydrochloride) (PAH) and PAH/poly(acrylic acid) (PAA) multilayers formedat the LC-aqueous interface were investigated by covalently attachingfluorescent dyes to both PSS and PAH, and by comparing the growthcharacteristics on LCs to solids with hydrophilic and hydrophilicsurfaces as a function of pH. Whereas the PEMs formed from PSS/PAH growat the interfaces of LCs in a manner comparable to the growth on thesurfaces of solids, the growth characteristics of PAA/PAH PEMs on LCsdiffer substantially from the solids investigated. While PAH/PAA filmsshow no growth on hydrophobic OTS surfaces their growth at theLC-aqueous interface was much higher than that observed on thehydrophilic solid surfaces. The zeta-potential measurements ofLC-aqueous interface also provides evidence for the presence of a smallnegative charge at these interfaces.

The inventors characterized the growth behavior of PSS/PAH multilayersat the 5CB-aqueous interface with the incorporation of fluorescentmolecules in both cationic and anionic polyelectrolyte. The presentstudy incorporated both FITC labeled PAH and Rhodamine labeled PSS withthe aim of investigating the growth behavior of PSS/PAH films at theLC-aqueous interface and to provide evidences for PSS adsorption at theLC interface.

The present example illustrates the characteristic features of PEMsgrown at the 5CB-aqueous interfaces and compares them to PEMs grown onsolid substrates. In addition, a PEM system of PAH/poly (acrylic acid)(PAA) films formed at the 5CB-aqueous interface is described. UnlikePSS/PAH multilayer system, PAH/PAA films consist of both weakpolyelectrolytes and hence shows a strong dependence on the pH of thepolyelectrolyte solutions. These films formed at a pH combination ofPAA(3.5)/PAH(7.5) are known to form very thick films and undergo a pHinduced reorganization. Such properties of these films make them usefulfor the control of the interfacial properties of the LC interface.

The growth of the PSS/PAH and PAH/PAA multilayer systems at the5CB-aqueous interface as compared to that on the hydrophobic andhydrophilic solid surfaces led to the identification of differences ingrowth behavior of these multilayer systems. While PSS/PAH films grow atthe 5CB-aqueous interface in a manner comparable to hydrophobic andhydrophilic solid substrates, PAH/PAA films grow at a higher rate at theLC-aqueous interface as compared to hydrophilic solid substrates and donot grow on hydrophobic substrates. This example lastly describes thenature of the LC-aqueous interface for being charged or hydrophobic innature by performing zeta-potential measurements of the LC in wateremulsions.

PEM formation at the LC-aqueous interface not only facilitates the useof this system for biological sensing by providing a functional andresponsive substrate unlike a solid substrate but also for the study ofother events such as pH or salt induced molecular rearrangements in thepolyelectrolyte film or the diffusion of molecules known to causeordering transition of LCs through PEMs formed by variouspolyelectrolyte systems. LC-aqueous interfaces provide a unique platformfor PEM growth due to their mobile nature on which various events suchas reorganization and molecular diffusion are expected to take placemore selectively.

FIGS. 7A and 7B show the experimental set-up and materials used in thisstudy respectively. The goal of the experiment reported in FIG. 8 was todetermine the growth characteristics of PEMs of PAH/PSS formed at theinterface between an aqueous phase and 5CB. In contrast to experimentswhere the pH of the aqueous electrolyte containing PAH was notcontrolled, the experimental results reported in FIG. 8 were obtained atpH 8. The results reported in FIG. 8 use FITC-PAH with PSS, and PAH withRh-PSS. In each of the experiments reported in FIG. 8, the interface wasfirst contacted with PSS or Rh-PSS. While the fluorescence is measuredafter the deposition of PSS-Rh in FIG. 8A, it is measured after thedeposition of FITC-PAH in FIG. 8B. For all the experimental measurementsdescribed in this example the error bars are calculated as the standarddeviation obtained from 2 to 3 independent measurements with multiplesamples at a time.

Inspection of FIG. 8 reveals that the growth in fluorescence ofrhodamine in FIG. 8A and FITC in FIG. 8B is linear with number of layersdeposited at the interface. The absolute values of fluorescenceintensity can not be compared between FIGS. 8A and 8B because themeasurements were performed with different fluorophores as well asimaging conditions.

Whereas the rate of growth of fluorescence from FITC-PAH in FIG. 3appears to decrease after 8^(th) bilayer, such a decrease in rate ofgrowth of the PEM is not seen in FIG. 8 b. The experiments reportedherein were performed using PAH solutions adjusted to pH=8, whereas thesolutions used in the previous experiments shown in FIG. 3 wereperformed using aqueous solutions of PAH that were not adjusted to pH 8.

The increase in fluorescent intensity relative to the background afterthe exposure of PSS-Rh to the 5CB-aqueous interface indicates theadsorption of PSS-Rh at the 5CB-aqueous interface. Also it should benoted that the increment in fluorescence intensity associated with thedeposition of the first layer of PSS-Rh was greater than the incrementin fluorescence intensity associated with deposition of subsequentlayers of Rh-PSS. This result suggested that the amount of Rh-PSSadsorbed onto the interface of the 5CB (first layer) is greater than theamount of Rh-PSS deposited onto the PEM. In contrast, the increment influorescence upon deposition of the first layer of FITC-PAH is similarto subsequent layers. The first layer of FITC-PAH is not adsorbeddirectly onto the aqueous interface of the 5CB but is adsorbed onto aPSS-decorated interface of 5CB.

The goal of the experiments shown in FIG. 9 was to compare the growthcharacteristics of PSS/PAH multilayers at the 5CB-aqueous interface withgrowth of PSS/PAH multilayers prepared on hydrophobic and hydrophilicsolid substrates. The multilayers were grown on OTS-treated glass andclean glass surfaces using the same set of polyelectrolyte and solutionconditions used to obtain the data in FIG. 8. The starting layer for the10 bilayer system was PSS for all experiments.

FIG. 9 shows the increase in fluorescence for the PSS/FITC-PAH filmsgrown at the interface of 5CB, clean glass and OTS-treated glass. FromFIG. 9, it is observed that: (a) a linear growth of fluorescence is seenat all three interfaces; (b) the rate of growth of fluorescence issimilar on all three interfaces, although the rate is slightly higher atthe 5CB-aqueous interface than OTS-treated glass; (c) independentellipsometric measurements (see FIG. 13) show that growth of the PEMs isslightly higher on native oxide surfaces than OTS-treated surfaces whichis consistent with the observations obtained from fluorescencemeasurements (on the right side of FIG. 9, the inventors have plottedthe ellipsometric thickness of the PEM obtained at the native oxidesurface). Because the growth of fluorescence for all the PEMs in FIG. 9is similar, the ellipsometric thicknesses obtained at the native oxidesurface can be used to estimate the thicknesses of the PEMs of PSS andPAH formed at the aqueous-LC interface. These results suggest that PEMsof PSS and PAH with thicknesses between 5 nm and 50 nm can be formed ataqueous-LC interfaces.

The experiments reported in FIG. 10 sought to characterize the effect ofthe pH of the weak polyelectrolyte PAH on the growth of PSS/PAH PEMs atthe 5CB-aqueous interface. As previously-reported, the pH of an aqueoussolution of the weak polyelectrolyte PAH influences the growth ofPSS/PAH multilayers on the surfaces of solids. At pH below 7, PAH insolution is largely protonated, whereas at pH's higher than 7 it becomespartially charged with its charge getting reduced to 88% of the maximumcharge at a pH of 8 in PEM environment (with no salt used). To testwhether a 5CB-aqueous interface behaves similar to a solid substrate,showing similar growth trends at different pH, the inventors chose a pHof 5 (fully charged) and 8 (partially charged) for FITC-PAH solution forPSS/FITC-PAH multilayer formation. In the multilayer formation at the5CB-aqueous interface, the weak polyelectrolyte solutions of FITC-PAHwas maintained at a pH of either 5 or 8, while the pH of strongpolyelectrolyte, PSS (pH=5.58) was not adjusted. The base layer for thedifferent substrates being the same as described above. It is importantto note that although the fluorescence of FITC-PAH is known to vary withpH, all fluorescence measurements were performed immediately afterrinsing the PEMs for 5 min at the pH (5.5-6.5) of rinsing water.

FIG. 10 shows measurements of the growth of fluorescence of PEMsprepared from FITC-PAH and PSS at the interface of 5CB at pH 5 or 8.From FIG. 10, it is observed that: the rate of growth of fluorescence isgreater at pH 8 than pH 5, consistent with prior reports of the effectof pH on growth of this PEM system. As shown in FIG. 14 the inventorsalso used ellipsometry to verify that the effect of pH on PEM growth onnative oxide and OTS-treated surfaces are similar. As shown in FIG. 10,the magnitude of the effect of pH on the growth of the PEM at theinterface of the 5CB is different from the effect of pH on the growth ofthe same PEM at the interface of OTS-treated glass. Not only the effectof pH is smaller but also unlike OTS coated glass the difference is onlyobserved after 8-9 bilayers in the case of multilayers grown at the5CB-aqueous interface.

The results in FIG. 10 demonstrate that the pH of the PAH solution doesnot influence the growth behavior of PSS/FITC-PAH multilayer films atthe LC-aqueous interface, thought the presence of a small effect may notbe reflected because of the large error bars associated with themeasurements with LCs. With a controlled pH of 5 and 8 of the FITC-PAHsolution, a linear growth trend is observed for the multilayers formedat the 5CB-aqueous interface. The multilayers were observed to bethicker for a PAH solution pH of 8 than a pH of 5 for all the substratesstudied.

The experiment reported in FIG. 11 sought to characterize growth ofFITC-PAH/PAA multilayers formed at the 5CB-aqueous interface. The growthof weak polyelectrolyte multilayers of FITC-PAH/PAA on a solid substrateis known to be highly pH dependent. With variation in pH, the chargedensity of the polyelectrolyte changes and hence different thicknessesof films are obtained after layer by layer deposition. Similar to theabove investigations for PSS/PAH multilayer films formed at the5CB-aqueous interface, here the inventors first established the growthtrend for FITC-PAH/PAA multilayer films formed at the 5CB-aqueousinterface. Second, the inventors compared the growth of FITC-PAH/PAAmultilayers formed at the 5CB-aqueous interface with those formed onsolid substrates; and third, The effect of two different pH combinations& polyelectrolyte concentrations on the multilayer film growth wasinvestigated.

In the process of multilayer formation, the 5CB-aqueous interface wasgently exposed first to FITC-PAH (1 mg/ml or 0.2 mg/ml, pH=7.5 or 6.5)for 15 min and then subsequently exposing the interface to poly acrylicacid (PAA, 1 mg/ml, pH=3.5 or 6.5) for 15 min after rinsing and thissequential procedure was repeated for 10 bilayers. To investigate theFITC-PAH/PAA PEMs formation on 5CB, two different pH systems werechosen, (a) FITC-PAH(7.5)/PAA(3.5)—which is known to show a significantpH induced thickness transition and (b) FITC-PAH(6.5)/PAA(6.5)—whichforms very thin layers because of the fully charged state of thepolyelectrolyte. To further investigate the nature of the growth ofPAH/PAA-multilayers grown at the 5CB-aqueous interface as compared tosolid substrates, multilayers were grown on hydrophilic native oxideglass and hydrophobic OTS coated glass with similar set ofpolyelectrolyte solutions as used for 5CB-aqueous interface.

FIG. 11 shows the increase in fluorescence for the FITC-PAH/PAA filmsgrown at the interface of 5CB, clean glass and OTS-treated glass for twodifferent pH systems. In all the cases FITC-PAH is the starting layer.It is evident from FIG. 11 that PAH/PAA multilayers grow for differentpH combinations of polyelectrolyte at the 5CB-aqueous interface.Inspection of FIG. 11 (for data corresponding to 5CB-aqueous interface)reveals an approximately linear growth of fluorescence for 7.5/3.5system while for 6.5/6.5 system the fluorescence increases up to 7^(th)bilayer and then remains almost constant. FIG. 11 also provides with theevidence of PAH adsorption at the 5CB-aqueous interface.

It can be observed from FIG. 11 that PAH/PAA multilayers grown atdifferent pH combinations show different rate of growth of fluorescenceat the interface of 5CB. Inspection of FIG. 11 reveals that the rate ofgrowth of fluorescence (for data corresponding to 5CB-aqueous interface)at pH 6.6/6.5 is less than pH 7.5/3.5. This result is qualitativelyconsistent with past reports of the effect of pH on the growth of thesePEMs. (FIG. 15 shows ellipsometric measurements of the effect of pH onthe growth of this PEM system on a native oxide surface under the sameconditions used to obtain data for the multilayers grown at the5CB-aqueous interface.) Inspection of growth pattern of FITC-PAH/PAAmultilayers on a native oxide surface reveals a qualitative trend thatis similar to that obtained for the 5CB-aqueous interface, in that theellipsometric thickness of the PEM deposited at pH 7.5/3.5 is greaterthan pH 6.6/6.5.

Inspection of FIG. 11 (for data corresponding to 5CB-aqueous interfaceand native oxide glass) reveals that rate of increase of fluorescencefor multilayers achieved in the case of 5CB-aqueous interface is muchhigher than that obtained on native oxide surface. This indicates thatthe incorporation of fluorescent polyelectrolyte is more at the5CB-aqueous interface than the native oxide surface. Closer inspectionof FIG. 11 also reveals that the ratio of fluorescence intensitiesobtained using PEMs deposited at pH 7.5/3.5 versus pH 6.5/6.5 atinterfaces of glass is larger than for the interface of 5CB.

FIG. 11 (for data corresponding to OTS coated glass) further illustratesthe growth behavior of PAH/PAA films on the OTS coated hydrophobic glasssurface. These measurements reveal that these PEMs do not grow onOTS-treated glass, emphasizing that the growth of these PEMs at theinterfaces of 5CB is significantly different from solid, hydrophobicsurfaces. The inventors observed the formation of an initial layer ofFITC-PAH on the OTS coated glass but layers do not deposit further toform multilayers. This result is in line with previously reportedresults. The adsorption of PAH on the hydrophobic surface of OTS coatedglass can be attributed to the negative zetapotential of OTS coatedsurface (reported in literature) or adsorption of hydrophobicpolyelectrolyte PAH in the hydrophobic environment of OTS SAMS.

To further compare the growth characteristics of PEMs on interfaces of5CB to surfaces investigated in the past, the inventors report in FIG.11 and FIG. 16 the effect of concentration of the polyelectrolytes (0.2mg/ml to 1 mg/ml). These experiments reveal the effect of concentrationof the polyelectrolytes on the growth of the PEMs at pH 7.5/3.5 appearsto be qualitatively similar but a little greater on the interface of 5CB(FIG. 1) than native oxide (FIG. 15) or glass (FIG. 11 and FIG. 16)surfaces.

In summary, PAH/PAA multilayers do grow at the 5CB-aqueous interface andunlike PSS/PAH PEMs, PAH/PAA multilayer films show different growthcharacteristic at the 5CB-aqueous interface as compared to solidsurfaces. While PSS/PAH multilayers grow at the 5CB-aqueous interface ina manner comparable to hydrophobic and hydrophilic solid substrates,PAH/PAA multilayer show significant differences. PAH/PAA multilayers atthe 5CB-aqueous interface grow at a rate which is much higher ascompared to the growth on the hydrophilic native oxide glass surface butthese multilayers simply do not grow on the hydrophobic OTS coated glasssurface for the mentioned solution conditions.

The results above indicate that the growth of PEMs on aqueous interfacesof 5CB differ from both native oxide/glass and hydrophobic surfaces.Perhaps most striking is the observation that PEMs of PAA and PAH growat interfaces of 5CB under conditions for which they do not grow athydrophobic surfaces (FIG. 11). It is also interesting that the resultsabove suggest that PAH adsorbs directly to the interface of the 5CB.

Inspection of FIG. 12 reveals that the zeta potential of the interfaceof 5CB at pHs above 5 is negative. The origin of the negative zetapotential is likely because of the strong adsorption of the hydroxylions at the 5CB-aqueous interface. The fact is further supported by thevariation of zetapotential of 5CB-aqueous interface with pH as shown inFIG. 12. This observation likely underlies the conclusion above that PAHadsorbs directly to 5CB (via electrostatic interactions and via theadsorption of hydrophobic polycation PAH in the hydrophobic environmentof 5CB). The results disclosed herein demonstrate that PSS/PAH andPAH/PAA multilayer films can be formed at the mobile 5CB-aqueousinterface.

For the case of PSS/PAH multilayers, the comparison reveals that the5CB-aqueous interface shows similar trends for the multilayer growth asshown by solid substrates. Although the PAH/PAA multilayers reveals agrowth trend at the LC-aqueous interface that resembles hydrophilicsolid but it differs significantly from solid hydrophobic surfaces.While PAH/PAA multilayer films do not grow on hydrophobic OTS coatedsurfaces they grow significantly at the 5CB-aqueous interface unlike thecase of PSS/PAH multilayers. The growth of PAH/PAA films is also muchhigher at the 5CB-aqueous interface as compared to hydrophilic solidsubstrates. Also, various factors like polyelectrolyte pH and solutionconcentration affect the film growth in a slightly different manner asthey do for solid substrates.

Zeta-potential measurements performed on the 5CB-aqueous interfacesindicate the presence of small negative charge at these interfaces (FIG.12) at pH values greater than about 5. The results thus obtainedprovides a firm ground for the future research which is focused on usingLCs in conjunction with PEMs to serve as amplifiers and transducers forvarious molecular events, thus providing approaches to the design of arange of materials with potential technological applications in highthroughput screening, drug delivery, and chemical and biologicalsensing.

EXAMPLE 10 Formation of Biomolecule-Containing PEMs at Aqueous-LCInterfaces to Report the Presence and/or Activity of Biomolecules

The coupling between LCs and PEMs that incorporate biomolecules providesopportunities to report the presence and activity of biomolecules. Forexample, the coupling can be used to: 1) observe the transport of activeenzymes through PEM films; 2) monitor the digestive activity of enzymeson PEM films that incorporate proteins; and 3) tailor PEM films tofacilitate detection of the presence and activity of protein toxins suchas botulinum toxin in solution.

Transport of Enzymes to Substrates Hosted at Interfaces of LCs.Phosphlipase A2 (PLA₂) and DLPC may be used as a model system todemonstrate that PEM films can mediate the transport of enzymes fromsolution to the aqueous-LC interface. First, PEM films are prepared fromPSS and PAH at aqueous-5CB interfaces that are decorated with DLPC (FIG.17). DLPC is dilaurylphosphatidylcholine. The number of layers of PSSand PAH is varied from 0-10 in order to determine the number of layersthat will prevent transport of the PLA₂ to the interface. The arrival ofthe enzyme at the DLPC-decorated interface of the LC is reported by anorientational transition associated with the enzymatic hydrolysis of theDLPC.

Past studies demonstrate that the permeability of proteins (IgGs)through PEMs can be increased through the incorporation of proteins intothe PEMs. PEMs prepared from PSS and IgGs may be used to show that theincorporation of IgG into the PEM permit permeation of the PLA₂ to theinterface under conditions for which PEMs formed from PSS and PAH do notpermit permeation. Again, the orientational ordering of the LCs isreported by the arrival of active enzyme at the interface.

PAA-PAH PEM films that are known to reorganize in response to changes inionic strength. These films can be used to control the transport ofenzymes to the aqueous-LC interface. DLPC can be deposited at theaqueous-5CB interface and then PEM films prepared from PAA and PAH on5CB. The films are incubated at high ionic strength in order tointroduce porosity. The films can be exposed to solutions of PLA₂ andchanges in the appearance of the 5CB monitored to determine the extentof transport of the PLA₂ across the porous film.

Protein-containing PEM Films as Reporters of Enzyme Activity. Whereasthe above paragraphs address the transport of proteins through PEMs byusing anchoring transitions of the LCs, PEMs which host enzymes and areformed on the surfaces of LCs offer opportunity to create materials forbiological sensors. Such materials are illustrated through descriptionsof examples that involve incorporation of PLA₂ into PEMs formed on LCs.

Past studies demonstrate that it is possible to incorporate proteinsinto PEMs. As depicted in FIG. 18, phospholipid substrates for PLA₂ thatare associated into vesicles will not permeate the PEMs (due to theirsize) whereas the products of enzymatic degradation will dissociate fromthe vesicles and form aggregates that are sufficiently small to be ableto permeate the PEMs. This difference in permeability of the lipidsmakes it possible to report the presence of the products of theenzymatic reaction by their permeation through the PEM and associatedorientational transition of the LC. The location of the enzyme withinthe PEM can be varied from the outer-most layer (highest accessibilityto vesicles) to 1-6 layers beneath the outermost layer.

Reporting Activity of Botulinum Toxin. This example describes the use ofPEMs formed on LCs for detection of botulinum neurotoxins (BoNTs). BoNTscause paralysis by cleaving proteins involved in acetylcholine releaseand are among the most potent of biological toxins. This example uses a19 amino-acid peptide sequence from SNAP-25, a synaptosomal protein thathas been identified as a substrate for BoNT/A. Experiments are performedusing the light chain of BoNT/A, which catalyzes the cleavage ofSNAP-25. As shown in FIG. 19, (i) double-tailed peptide-amphiphiles fromthe 19 amino acid sequence of SNAP-25 is incorporated into vesicles ofDLPC that do not permeate a PEM (e.g., PSS/PAH) formed at the surface ofa LC, (ii) the light chain of BoNT/A is introduced into the aqueousphase, and (iii) cleavage of the peptide-amphiphile leads to itstransport through the PEM and thus is reported by the liquid crystal.

EXAMPLE 11 Formation of PEMs of Hyaluranon (HA) and Collagen (COL) onLCs

This example describes the formation of PEMs from COL and HA on theinterfaces of LCs. HA, also called hyaluronic acid, is a polysaccharideextracellular matrix (ECM) molecule abundant in tissues, and tightregulation on its synthesis controls cell adhesion and motility duringcancer metastasis, stem cell homing, wound healing, etc. Collagen is apredominant ECM protein as well. To prepare PEMs of (HA/COL) of order“n”, with n representing the number of adsorbed layer pairs, HA and COLsolutions are prepared as follows. Collagen of 1 mg/ml in acetic acid(Sigma, St. Louis, Mo.) is diluted to 0.2 mg/ml with water. Hyaluronan(MW=400 KDa, Barcelona, Spain) is dissolved in pure water at 1 mg/ml.Both solutions are adjusted to pH 4 with HCl. Between each adsorptionstep, the films are extensively rinsed with 10⁻⁴M HCl. To characterizegrowth of the PEM, the collagen stock is dialyzed to concentrate thecollagen. Dialysis removes salts that may interfere with the labelingreaction. The concentrated collagen is adjusted to 8 mg/ml and pH 8.3for the labeling reaction with Alexa430-conjugated succinimidyl ester.This ester reacts with unprotonated amine. The ester solution isprepared in anhydrous DMSO with a 1:20 ratio to amount of collagen. Freeester is hydrolyzed in about 24 hours. The reaction mixture is incubatedat room temperature for an hour before it is ready to be diluted anddeposited on HA. Fluorescence intensity is approximately proportional toPEM thickness, a relationship shown in previously published data.

EXAMPLE 12 PEMs Formed on LCs can be Used to Report Specific BindingEvents Between Proteins

PEMs of HA and COL are prepared on LCs, as described in the examplesabove. The optical appearance of the LC is determined by using polarizedlight microscopy and a birefringence mapper. Following formation of thePEMs, anti-collagen antibodies (1 micromolar) at added to the aqueousphases. A change in optical appearance of the LC reports the presence ofthe antibodies. In contrast, when anti-biotin antibodies are introducedinto the aqueous phase there is no change in optical appearance of theLC. This example serves to illustrate the way in which PEMs formed onLCs can be used to report specific binding events between proteins.

EXAMPLE 13 Formation of PEMs Containing Biomolecules on LC EmulsionDroplets

PEMs on LC droplets are useful systems for drug delivery and providebiophysical models of cells. In this example, LC emulsions (i.e.,particles made of LCs dispersed in water) will be used as templates forthe deposition of PEMs. A distinct advantage offered by emulsions is (i)their large surface area, potentially offering orders of magnitudegreater sensitivity for recording molecular interactions with the LCinterface; and (ii) the prospect of forming mobile and passive opticalreporters of the presence of targeted chemical and biological species,and (iii) the opportunity to further engineer these systems through thedistortions and defects formed by LCs in confined, spherical geometries.Such materials might find use, for example, as “smart colloids” capableof reporting biological activity (e.g., BoNT/A) when added to a sampleor incorporated into a coating.

In this example, PEMs of HA and COL are prepared on LC emulsiondroplets, as described in the examples above. The optical appearance ofthe LC is determined by using polarized light microscopy and abirefringence mapper. Following formation of the PEMs, anti-collagenantibodies (1 micromolar) at added to the aqueous phases. A change inoptical appearance of the LC reports the presence of the antibodies. Incontrast, when anti-biotin antibodies are introduced into the aqueousphase there is no change in optical appearance of the LC emulsiondroplets. This example serves to illustrate the way in which PEMs formedon LC emulsion droplets can be used to report specific binding eventsbetween proteins.

EXAMPLE 14 PEM Coating of Emulsions

In this example, the formation of polyelectrolyte multilayers onthermotropic LC (oil)-in-water emulsions is described. The growth of thepoly(styrene sulfonate) (PSS)/poly(allylamine hydrochloride) (PAH)multilayers was characterized with microelectrophoresis, flow cytometry,and fluorescence microscopy. Up to eight PSS/PAH bilayers were depositedand the effect of the multilayer formation on the orientation of the LCwas examined using polarized light microscopy. The inventors alsoexamined the influence of the multilayer coating on the interactionbetween an analyte (surfactant) and the LC core. The PSS/PAH-coated LCdroplets were subsequently treated with ethanol to dissolve the LC coresto form hollow capsules. The production of stable capsules confirmed thegrowth of the multilayer films, and the integrity of the film formed.These results are significant in that (a) they demonstrate the abilityto prepare stable LC emulsions with an easily modifiable interface, and(b) they show that LC emulsions can be used to template polyelectrolytemultilayer formation. These developments are significant because of thepotential benefits in sensing and encapsulation applications.

Emulsion Preparation. The LC-in-water emulsions were formed bysonicating a mixture of 1 vol. % 5CB in a dispersant at 10 W for 60 s.Three types of dispersants were used: (a) water, (b) an aqueous solutionof a strongly charged polyelectrolyte, poly(styrene sulfonate) (PSS) (1mg mL⁻¹), and (c) an aqueous solution of a cationic phospholipid,1,2-dilauroyl-sn-glycero-3-ethylphosphocholine (DLEPC) (700 μM). Theresulting emulsions were denoted as 5CB-H₂O, 5CB-PSS and 5CB-DLEPC,respectively. The LC droplets in all three types of emulsions werespherical, with a size range of 1-10 μm and were visually observed to bestable against coalescence for months. The 5CB-PSS and 5CB-DLEPCemulsions are stable as PSS and DLEPC can associate with the interfaceof the water-immiscible droplets of 5CB, effectively acting likesurfactants, electrostatically stabilizing the droplets.

The ζ-potentials of 5CB-H₂O, 5CB-PSS and 5CB-DLEPC droplets were −40 mV,−45 mV and +40 mV as prepared. (measured in water at pH ˜5.7). Theζ-potentials were also measured over a wide pH range to determine theisoelectric point (IEP) of the droplets. The IEP for the 5CB-H₂Odroplets was found to be 5, and the IEP shifted to 3.5 and 7.0 for the5CB-PSS and 5CB-DLEPC droplets, respectively (FIG. 21 a). This isindicative of the PSS and DLEPC providing charge stabilization for theemulsions. Notably, 5CB did not form a stable emulsion in water when PAHwas used instead of PSS. This result is similar to previous findings;PSS can render uncharged pyrene water-dispersible, but PAH does not.This result suggests that successful emulsification of 5CB with PSS maybe attributed to the amphiphilic nature of PSS (the aromatic group ishydrophobic while the charged sulfonate group is hydrophilic) and/or theπ-π interactions between the phenyl groups of 5CB and PSS.

The stability of the 5CB-H₂O emulsion is likely due to the spontaneousadsorption of hydroxide ions at the oil-water interface, which has beenreported for surfactant-free emulsions of linear alkanes such ashexadecane and dodecane. In those studies, the oil droplets werenegatively charged and the magnitude of their ζ-potential depended onthe pH and the ionic strength of the aqueous phase. This phenomenon hasbeen reported to be specific to the hydroxide ion; other cations andanions are not preferentially adsorbed at the oil-water interface. Thisspecificity has led to the suggestion that adsorption of the hydroxideion may involve hydrogen bonding with the water molecules at theinterface. The 5CB-H₂O emulsion exhibits a similar behavior; it isnegatively charged as prepared (−40 mV, measured in water at pH ˜5.7),and this charge is pH-dependent. The IEP the 5CB-H₂O emulsion is 5,which is slightly higher than those reported for oil-in-water emulsionsof linear alkanes (IEP ca. 3-4).

Multilayer Coating of Emulsions. The polyelectrolytes for LbL assemblywere deposited from 1 mg mL⁻¹ solutions containing 0.1 M NaCl. Theelectrolyte is added to facilitate polyelectrolyte adsorption. In theabsence of salt, the polyelectrolytes adopt a stretched conformation andadsorb as thinner, more rigid layers, which might result in incompleteand/or less homogeneous coverage of the LC droplets. The pH of the PAHsolution was adjusted to pH 7 (pK_(a) of PAH is ˜8.5) while the pH ofthe PSS solution was not adjusted (PSS is a strong polyelectrolyte). The5CB-H₂O emulsions were negatively charged as prepared; hence thecationic PAH was deposited as the first layer. Despite the stability ofthe 5CB-H₂O emulsions, the LbL assembly with PAH and PSS did not proceedas well as with the 5CB-DLEPC or 5CB-PSS emulsions. ζ-potentialmeasurements of these samples show inconsistent magnitudes and a broaddistribution of charges (data not shown). Flow cytometry was also usedto quantify the fluorescence intensity of the 5CB-H₂O emulsion dropletsafter alternating adsorption of fluorescein isothiocyanate-labeled PAH(PAH-FITC) and PSS. Although there was a general increase influorescence intensity over 14 adsorption steps, the intensity was lessthan half of that observed with 5CB-PSS and 5CB-DLEPC emulsions. Theinventors also observed desorption of PAH-FITC after coating; thefluorescence measured in the aqueous phase of 5CB-H₂O emulsions with aPAH-FITC terminated layer slowly increased after the adsorption andwashing steps. These results indicate that PAH-FITC was weakly bound tothe 5CB-H₂O core, resulting in incomplete polymer coverage of thedroplets and the surface of these droplets having both positive andnegative charges, accounting for the poorer stability of theseemulsions.

On the other hand, the 5CB-PSS and 5CB-DLEPC emulsions were coatedsuccessfully with at least 16 layers of PAH and PSS. The alternatingζ-potentials for both the 5CB-PSS and 5CB-DLEPC emulsions during LbLassembly (between +40 mV and −50 mV) are consistent with stepwiseadsorption of cationic PAH and anionic PSS (FIG. 21 b). These values aresimilar to ζ-potentials of PSS and PAH alternately deposited on chargedsolid particles. As each layer is adsorbed, the surface charge isreversed, allowing the adsorption of the next layer via electrostaticinteractions.

However, ζ-potential measurements only serve as a qualitative indicationof multilayer growth. As such, PAH-FITC was used to further confirmmultilayer growth on 5CB-PSS and 5CB-DLEPC emulsions using flowcytometry. The increase in fluorescence intensity of the droplets wasquantified using flow cytometry (FIG. 22 a). The growth in fluorescenceintensity was linear after the third and first layer for 5CB-PSS and5CB-DLEPC emulsions, respectively. The thinner initial layers for the5CB-PSS system indicate that it takes a few deposition steps for thelayer thickness to reach an equilibrium value. This can be explained bysubstrate effects, which are commonly observed in the first few layersin LbL systems. A fluorescence image of 5CB-PSS coated with sevenbilayers of PAH-FITC/PSS shows uniform fluorescence around the droplets,confirming the flow cytometry results (FIG. 22 b). Similar images wereobtained for 5CB-DLEPC emulsions coated with seven bilayers ofPSS/PAH-FITC (data not shown). The results obtained frommicroelectrophoresis, flow cytometry, and fluorescence microscopy alldemonstrate that PE multilayer growth at the mobile interface of 5CBemulsion droplets is comparable to similar multilayers assembled onsolid, charged colloidal particles.

Orientation of 5CB in the Emulsions. Polarized microscopy was used tostudy the orientation of the 5CB at the droplet interface before andafter deposition of the multilayer films. Past studies have establishedthat the orientations of a LC within a droplet depend on factors such asthe bulk elasticity of the LC, the orientation of the easy axis of theLC at the interface of the droplet, and the anchoring energy of the LC.For sufficiently large droplets, surface anchoring dominates, whichresults in droplets that contain topological defects at equilibrium. Theinventors observed the orientation of 5CB in the emulsion droplets to beindependent of droplet size. This implies that the droplets are at thelimit of strong anchoring. Previous studies have also shown that planaranchoring of 5CB at the interface results in a bipolar configurationwhere each droplet contains two or more point defects at the interface,known as ‘boojums’. On the other hand, homeotropic anchoring of 5CB atthe interface results in a radial configuration, where each droplet hasa point defect at its center, known as a ‘hedgehog’. Optical images(crossed polars) of 5CB-H2O and 5CB-DLPC emulsion droplets are shown inFIG. 23 along with the orientations of the 5CB within the droplets. The5CB-H₂O emulsion droplets have a bipolar configuration with two pointdefects at the surface, while the 5CB-DLEPC emulsion droplets have aradial configuration, with characteristic cross-like appearances. Theappearance of the 5CB-PSS emulsion droplets was similar to the 5CB-H2Odroplets. The orientations of the LC at the interfaces of the dropletsare similar to the orientations of 5CB and DLEPC-decorated 5CB reportedpreviously at planar LC-aqueous interfaces. FIG. 24 shows optical images(crossed polars) of 5CB-PSS and 5CB-DLEPC emulsion droplets coated with5 bilayers of PAH/PSS. The results reveal that the bipolar and radialconfigurations of LC within the 5CB-PSS and 5CB-DLEPC emulsion dropletsare preserved during multilayer coating.

Effect of surfactants on the Orientation of 5CB in the Emulsions. Inprevious studies, the exposure of 5CB to surfactants such as sodiumdodecyl sulfate (SDS, anionic) caused a rapid ordering transition from aplanar to a homeotropic orientation of 5CB. Preliminary studies on theuncoated and (PAH/PSS)₇-coated emulsions showed that exposure to 5 mMSDS triggered an ordering transition of 5CB from a planar to ahomeotropic orientation. This was reflected by the appearance of crosseson the emulsion droplets when viewed with cross-polarizers (FIG. 25).The ordering of 5CB in the uncoated emulsions changed from planar tohomeotropic instantaneously, however this change took about five min tooccur for the (PAH/PSS)₇-coated emulsions. This result indicates thatSDS can permeate the multilayer through to the LC core. In addition, themultilayer can influence the way SDS penetrates and adsorbs at the 5CBinterface by slowing the diffusion kinetics.

Hollow Capsules. To further verify the structural integrity of themultilayer coating on LC emulsions, the inventors selectively removedthe LC core of the multilayer-coated emulsion droplets by dissolution ofthe LC with ethanol to yield hollow polyelectrolyte capsules. Theinventors found that hollow capsules could only be obtained frommultilayer-coated emulsions with a PSS terminated layer. Emulsionscoated with a PAH terminated layer aggregated upon treatment withethanol. It is possible that the ionization of PAH at the surfacedecreases (via conversion of the ammonium groups to amines) when exposedto ethanol, thereby destabilizing the emulsion. Decreasing solventpolarity (e.g. increasing ethanol content) of the PAH depositionsolutions can result in a dramatic increase of layer thickness as aconsequence of charge screening and the polyelectrolytes adopting a morecoiled conformation. The PSS-terminated multilayer-coated emulsionsturned clear immediately after the addition of ethanol. This is a commonindication of core dissolution in other multilayer systems where solidcore particles are used.

The hollow capsules were characterized with transmission electronmicroscopy (TEM) and atomic force microscopy (AFM). The capsule wallshave a grainy texture, which is probably due to slight rearrangement ofpolyelectrolytes when they were exposed to ethanol. When dried, thecapsules collapse and the folds are visible from TEM and AFM images(FIG. 26). AFM was used to obtain the thickness of the capsule walls bytaking a cross-sectional profile of the capsules where it was foldedonly once. Hence, the measured thickness corresponds to twice themultilayer wall thickness. The average thickness per layer wasdetermined by taking the average thickness of several cross-sectionalprofiles and dividing by twice the total number of layers deposited. Thelayer thickness was calculated to be approximately 1.4 nm for bothmultilayer-coated 5CB-PSS and 5CB-DLEPC emulsions, which correspondswell to typical layer thickness of the same polyelectrolyte multilayersassembled on planar and colloidal supports. This study also demonstratesthat thermotropic LCs, despite having a mobile interface, can be used ascores in core-shell systems. The ability to synthesize monodisperseemulsions at a desired size overcomes difficulties currently faced whenusing solid particles as templates of polyelectrolyte capsules. Forexample, it is often difficult to completely remove melamineformaldehyde (MF) cores; some residual MF may remain in the capsulewalls.

Experimental Section.

Materials. Poly(sodium-4-styrenesulfonate) (PSS, M_(w), 70 kDa),poly(allylamine hydrochloride) (PAH, M_(w), 70 kDa), and sodium dodecylsulfate (SDS) were purchased from Sigma-Aldrich and used without furtherpurification. Fluorescein isothiocyanate-labeled PAH (PAH-FITC) wasprepared as described previously. The nematic liquid crystal4′-pentyl-4-cyanobiphenyl (5CB) was purchased from EMD Chemicals(Hawthorne, N.Y.) and used without further purification. Thephospholipid 1,2-dilauroyl-sn-glycero-3-ethylphosphocholine (DLEPC) wasobtained from Avanti Polar Lipids (Alabaster, Ala.). An inline MilliporeRiOs/Origin system was used to produce high-purity water with aresistivity greater than 18.2 MΩ·cm.

Emulsion Preparation. The LC emulsions (1 vol. %) (typically 1 mL) wereformed by mixing 10 μL of 5CB with 1 mL of dispersant with a tipsonicator at a power of 10 W for 60 s. Three types of dispersants wereused: 1) water, 2) aqueous solution of PSS (1 mg mL⁻¹, no added salt),and 3) aqueous solution of DLEPC (700 μM).

Microelectrophoresis. ζ-potentials were measured using a MalvernZetasizer2000 instrument. Approximately 2 μL of 1 vol. % emulsions wasadded to 3.5 mL of water. The quoted values were calculated by takingthe average of 5 successive measurements. The pH of water was adjustedby adding NaOH or HCl no more than 5 min before measurement.

Multilayer Coating of Emulsions. 5CB-PSS and 5CB-DLEPC emulsions werefirst washed 3 times with water. Approximately 1 mL of emulsion and 1 mLof water was added to a 2 mL Eppendorf tube. The tube was agitated witha vortex mixer and then centrifuged at 5000 g for 5 min. This resultedin a pellet forming at the bottom of the tube. Approximately 1 mL of thesupernatant was removed and replaced with water. This was repeated twiceto remove excess PSS and DLEPC prior to polyelectrolyte coating. The PAHand PSS solutions for LbL were made to 1 mg mL⁻¹ containing 0.1 M NaCland PAH-FITC was made to 0.5 mg mL⁻¹ containing 0.1 M NaCl. Afterwashing, 1 ml of polyelectrolyte solution was added to 1 mL of theemulsion. The mixture was agitated with a vortex mixer and allowed toincubate for 15 min. After adsorption, the dispersions were centrifuged(at 5000 g, 5 min), after which the supernatant was removed and replacedby water. Washing was performed 3 times, followed by adsorption of thenext polyelectrolyte. The entire process was repeated until the desirednumber of layers was achieved. The initial 5CB-PSS emulsion isnegatively charged (as determined from ζ-potential measurement), so thefirst polyelectrolyte adsorbed is PAH. Conversely, since 5CB-DLEPCemulsion is positively charged, the first layer deposited was PSS.

Flow Cytometry. Fluorescence measurements of the (PAH-FITC/PSS)-coatedLC emulsions were carried out using a Becton Dickinson FACS Calibur flowcytometer. 1 μL of 1 vol. % emulsion was diluted with 200 μL of waterand this solution was analyzed on the instrument. Measurements wereacquired with triggering on the forward scatter detection (E0 detector)with a threshold of 400. FITC fluorescence was monitored on the FL1(515-545 nm) parameter with a PMT voltage of 550 V. For each sample,30,000 particles were analyzed at a rate of approximately 500 particlesper second. For each incubation time, a background sample was recorded(i.e., the same sample before any polyelectrolyte coating). This signalwas subtracted from the signal of the particles coated with a differentnumber of fluorescent layer. Flow cytometry data analysis was performedwith Summit v. 3.1 (Cytomation, Inc., Colorado, USA). A sub-populationwas analyzed by applying a gate within the entire population to excludeparticle aggregates and the same gate was used for one set of sampleswith different layer numbers. The mean fluorescence intensity wasobtained from the fluorescence intensity histograms.

Polarized and Fluorescence Microscopy. The orientation of the LC withindroplets was examined with plane-polarized light in transmission mode onan Olympus IX 71 inverted fluorescence microscope with crossedpolarizers. Fluorescence images were taken using the same microscopewith a FITC filter cube. In both cases, a 60× oil immersion objectivewas used and images were captured with a color camera.

Preparation of Hollow Capsules. Multilayer-coated 5CB-PSS and 5CB-DLEPCemulsions with an outermost PSS layer were treated with ethanol.Approximately 1 mL of ethanol was added to ˜1 mL of emulsion and themixture was agitated with a vortex mixer and allowed to stand for 15min. The mixture was then centrifuged, the supernatant removed and freshethanol was added. This was repeated three more times. After the lastethanol addition, the samples were allowed to stand overnight to ensurecomplete dissolution and removal of 5CB. Before characterization, thehollow capsules were washed three times with water. Air-dried hollowcapsules were characterized with a Philips CM120 BioTWIN transmissionelectron microscope (TEM) operated at 120 kV and a Nanoscope IIIa atomicforce microscope (AFM) (Digital Instruments Inc., Santa Barbara, Calif.)operated in Tapping Mode™ using silicon cantilevers with a resonancefrequency of ca. 290 kHz (MikroMasch, USA). Image processing(first-order flattening) was carried out with Nanoscope 4.43r8 software.

Those skilled in the art will recognize, or be able to ascertain usingno more than routine experimentation, numerous equivalents to thespecific materials and methods described herein. Such equivalents areconsidered to be within the scope of this invention and encompassed bythe following claims.

1. A method for providing a polyelectrolyte multilayer film at aliquid-liquid interface, comprising steps of sequentially-depositinglayers of anionic and cationic polyelectrolytes at a liquid-liquidinterface that is formed between immiscible first and second liquids,wherein the first liquid is an aqueous solution and the second liquid isa liquid crystal, whereby a polyelectrolyte multilayer film is providedat said liquid-liquid interface.
 2. The method according to claim 1wherein the liquid crystal is in the form of a droplet.
 3. The methodaccording to claim 1 wherein the second liquid is in the form of adroplet.
 4. A method for providing a polyelectrolyte multilayer film atan aqueous-liquid crystal interface, comprising steps ofsequentially-depositing layers of cationic and anionic polyelectrolytesat an aqueous-liquid crystal interface that is formed between an aqueousphase and a liquid crystal phase whereby a polyelectrolyte multilayerfilm is provided at said aqueous-liquid crystal interface.
 5. The methodaccording to claim 4 wherein the polyelectrolyte multilayer film iscapable of interacting with an analyte present in the aqueous solutionthereby causing a change in orientation or ordering of the liquidcrystal.
 6. The method according to claim 4 wherein the polyelectrolytemultilayer film includes an excipient capable of interacting with ananalyte present in the aqueous solution thereby causing a change inorientation or ordering of the liquid crystal.
 7. The method accordingto claim 6 wherein said excipient is a ligand or a receptor capable ofselectively-binding said analyte, an enzyme capable of catalyzing atransformation of the analyte, a molecule capable of undergoing achemical reaction in the presence of the analyte, or an enzyme substratecapable of being transformed by said analyte.
 8. The method according toclaim 4 wherein the polyelectrolyte multilayer film is depositeddirectly on the liquid crystal phase.
 9. A method for providing anunsupported polyelectrolyte multilayer film comprising steps of: (a)providing a liquid-liquid interface between immiscible first and secondliquids, wherein the first liquid is an aqueous solution and the secondliquid is a liquid crystal; (b) sequentially-depositing layers ofcationic and anionic polyelectrolytes at the liquid-liquid interfacewhereby a polyelectrolyte multilayer film is provided at theliquid-liquid interface; and (c) removing the first and second liquidsto provide an unsupported polyelectrolyte multilayer film.
 10. Themethod according to claim 9 wherein the second liquid is in the form ofa droplet.
 11. The method according to claim 9 wherein the unsupportedpolyelectrolyte multilayer film is in the form of a hollow capsule.